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ICCBM15
15th International Conference on the
Crystallization of Biological Macromolecules
September 17 - 20, 2014 – Hamburg, Germany
PROGRAM , ABST RACT S & GU I DELI N ES
visit www.ICCBM15.org
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Local Board
Christian Betzel, University of Hamburg
Jeroen R. Mesters, University of Luebeck
Sven Falke, University of Hamburg
Dominik Oberthür, CFEL/DESY and University of Hamburg
Workshop Directors
Edward H. Snell, Hauptman-Woodward Institute, Buffalo, USA
Jose A. Gavira, LEC (IACT), CISC-UGR, Granada, Spain
Christian Biertümpfel, MPI of Biochemistry, Martinsried, Germany
Scientific Board
Henry Chapman, CFEL/DESY and University of Hamburg, Germany
Yin Da-Chuan, NWPU, Xi‘An, PR China
Ursula Egner, Bayer Pharma AG, Berlin, Germany
Howard Einspahr, Editor, Acta Cryst, Section F, USA
John R. Helliwell, University of Manchester, UK
Michael Hennig, Hoffmann - La Roche, Basel, Switzerland
Rolf Hilgenfeld, University of Luebeck , Germany
Derek Logan, University of Lund, Sweden
Lawrence de Lucas, University of Alabama, Birmingham, USA
Thomas Marlowitz, CSSB/UKE/DESY, Hamburg &
IMP/IMBA, Vienna, Austria
Rob Meijers, EMBL Hamburg, Germany
Janet Newman, CSIRO, Parkville, Australia
Pavlina Řezáčová, )MG-ASCR, Prague, Czech Republic
Claude Sauter, CNRS IBMC, Strasbourg, France
Sujata Sharma, AIIMS, New Dehli, India
Katsuo Tsukamoto, Tohoku University, Sendai, Japan
Peter G. Vekilov, University of Houston, USA
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Hamburg, September 2014
conference venue: Edmund-Siemers-Allee 1, D-20146 Hamburg
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ICCBM15
Program overview
17. 09. 2014
08:00 – 08:55
08:55 – 09:20
Wednesday
Registration
Opening ceremony
Prof. C. Leopold, Prof. E. Hartmann, Prof. E. Weckert
09:25 – 10:05
Plenary/keynote: Leonid A. Sazanov
Membrane protein crystallization
10:40 – 12:30
Vadim Cherezov, Poul Nissen, Hartmut Luecke, Francoise Bonneté
Crystallization for lead compounds in industry
13:40 – 15:10
Jörg Benz, Linda Öster, Pamela Williams
Screening for crystals
15:50 – 17:20
Fabrice Gorrec, Simon Newstead, Edward Snell
18. 09. 2014
Thursday
08:15 – 08:55
Registration
Complementary methods
09:00 – 12:00
Trevor Forsyth, Anthony Watts, Georgeos Skiniotis, Jan-Pieter Abrahams, Henning Stahlberg
Nucleation: Theory and practise
13:00 – 14:35
Dominique Maes, Elias Vlieg, Jan Sedzik, Dimitri Radajewski
In-vivo crystallization
15:50– 17:05
Fasseli Coulibaly, Cait MacPhee, Daniel Passon
17:30 – 18:10
Keynote: Richard Giegé
20:00
Conference dinner
19. 09. 2014
Friday
08:15 – 08:55
Registration
Crystallization and scoring for SFX
09:00 – 10:40
Andrew Aquila, Petra Fromme, Guillermo Calero, Gergely Katona
Microfluidics crystallization, i.e. counter diffusion
11:20 – 12:35
Sachiko Takahashi, Sarah Perry, Michael Heymann
Crystallization for neutron diffraction
13:40 – 14:55
Zoë Fisher, Akihiko Nakamura, Tobias Schrader
Microgravity
15:45 – 17:00
Shigeru Sugiyama, Abel Moreno, Da-Chuan Yin
17:20 – 18:00
Keynote: Werner Kühlbrandt
20. 09. 2014
Saturday
08:30 – 08:55
Registration office
Scoring and optimization in crystallization
09:00 – 10:30
Janet Newman, Martin Caffrey, Bostjan Kobe
11:10 – 12:10
Vasundara Srinivasan, Alexandra Ros
12:15 – 12:55
Keynote and closing lecture: Christopher Tate
13:00 – 13:15
Discussion, farewell address, end of ICCBM15
Poster sessions: 17.09. (17:20 – 18:40), 18.09. (14:40 – 15:45),
19.09. (10:40 – 11:20, 12:45 – 13:40 and 15:00 – 15:40)
Exhibition: The exhibition is open during conference hours.
Speakers are asked to do a media check and contact the local staff in the lecture hall prior to their talk.
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WELCOME TO HAMBURG! – “Das Tor zur Welt”
Going back to ICCBM1 in 1985 this conference has taken place
approximately every two years in countries all over the world, including the
USA, France, Germany, Japan, Spain, China, Canada, Mexico and Ireland.
Hamburg is proud and honored to welcome you, hosting the 15th
conference in this long and successful streak, aiming to bring newcomers
and leading experts from different related fields together, dealing with the
crystallization of biological macromolecules. One scientific focus of this
event will be the application of novel radiation sources and innovative
crystallographic methods. Leisure time and organized tours, covering the
Best-of-Hamburg attractions such as Port of Hamburg, Fishmarket,
Jungfernstieg, Speicherstadt, Elbphilharmonie, Miniature Wonderland, St.
Pauli and Hagenbeck Zoo, might be exciting and entertaining for you as well.
“Dear colleagues, postdocs, students and friends,
It is our great pleasure to invite you to join ICCBM15 with focus on
macromolecular crystallization for novel radiation sources such as free
electron lasers and use of complementary methods (e.g. CryoEM). The
exciting program features many highly-esteemed colleagues from all over the
world.
For the individual desiring to learn how to grow crystals or to acquire new
crystallization skills, the conference is preceded by a 2.5 days workshop
consisting of short presentations, practical demonstrations and hands-on
exercises taught by an international team of experts.
We very much look forward to personally welcome you and/or your colleagues
in Hamburg for an illuminating meeting.”
Christian Betzel and Jeroen Mesters
Chairpersons
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Content
This is Hamburg: Map
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This is Hamburg: Around the conference building
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Public transport
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General information for your orientation
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Hotel contact
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Tourism information
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Some recommended restaurants
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Conference dinner
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DESY campus map
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How to get access to the internet
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Detailed scientific program
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Exhibition
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Oral presentations: Abstracts and speaker list
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Poster presentation abstracts
74
Index of authors
163
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ELBE
DESY
ALSTER
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ELBE
© open street maps and contributors
"Hamburg SPOT 1633L" by Cnes - Spot Image - http://gallery.spotimage.com/product_info.php?products_id=1633.
Licensed under Creative Commons Attribution-Share Alike 3.0 via Wikimedia Commons http://commons.wikimedia.org/wiki/File:Hamburg_SPOT_1633L.jpg#mediaviewer/File:Hamburg_SPOT_1633L.jpg
For your orientation in Hamburg
http://english.hamburg.de/tourist-information/
might be helpful.
Hamburg from high above… the enframed area is
shown above in more detail with the Hamburg
airport located in the north.
HARBOUR
Central Train Station
Conference Building
University of Hamburg
AIRPORT
This is Hamburg: Map
The following maps should provide a first insight into location and characteristics of Hamburg and its
districts. It is a “free” and “hanseatic” city – the eighth largest in the European union. Going back in history
it was a part of the hanseatic league a powerful trading and defensive confederation in northern Europe
between the 13th and 17th century. There is a very large precious harbor and for instance the International
Tribunal for the Law of the Sea is located Hamburg.
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Enjoy the colorful
gardens and entertaining
events in this park
“Planten un Blomen”
• Gas station, newspapers, fast food etc.
• Rent a bike (“StadtRAD”)
• Public transport and Taxi
© open street maps and contributors
Follow this road to cross the ALSTER (e.g.
to get to central station) or go downtown
shopping, relaxing, sight seeing…there are
also some restaurants as well as a cinema
not far away
Conference venue,
Edmund-Siemers Allee 1
University of Hamburg
surrounded by park areas
Dammtor station (arrival by train/”S-Bahn”,
line S11, S21 and S31)
entrance
Along Grindelallee you find a large
variety of small shops and
restaurants around the university
buildings
ALSTER
↑
N
This is Hamburg: Around the conference building
DESY
"Bahnlinien im HVV" by Lars Hänisch - Own work. Licensed
under Creative Commons Attribution-Share Alike 3.0 via
Wikimedia Commons http://commons.wikimedia.org/wiki/File:Bahnlinien_im_HVV.png
#mediaviewer/File:Bahnlinien_im_HVV.png
(easily reached by bus)
airport
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Central station (regional trains)
(station: „Landungsbrücken“) – look for
the green ship „Rickmer Rickmers“
Conference dinner
Conference venue – „Dammtor“
Public transport in Hamburg –
Train Railway lines („U-Bahn“ and „S-Bahn“ lines)
For information on the local bus lines, e.g. „HVV Metrobus“ line 4 or line 5 next to the
conference venue to travel to the city centre, ask the local conference staff or have a look
online. Plans and details and prices of special tickets are also displayed at every station and
online (www. hvv.de).
General information for your orientation – part 1
• All lectures will take place at the “main building” of the University of Hamburg (second
floor, lecture hall B), located in Edmund-Siemers-Allee 1 (picture, see map) close to the
station “Dammtor” as you see on the previous maps in this booklet. By the way the
University of Hamburg was founded in 1919. At the moment more than 40.000 students
are educated here, a large variety of university courses is offered.
• Local coordinators among the large organisation team:
Constanze Weismantel
General event management
Friederike Meyn
General event management
Sven Falke
IT and lecture hall technique, posters
Madeleine Künz
Catering, reception, transport
• The Conference fees include:
- Assistance/advice with travel- and hotel-issues
- Attendance of all conference and poster sessions
- Coffee breaks and lunches during the conference
- Conference bag with abstract book and further material
- Certificate of participation / presentation (upon request)
• The registration and information desk is open starting from 8 am on September 17th for the whole
conference. It is located on the ground floor of the venue. Maps of the venue floors are part of this booklet
(see „Exhibition“ and „Poster presentations“)
• To get access to WLAN you need an individual password from the local registration desk. More
Information is found later on in this booklet and available at the registration desk.
• Lost and found again: If you find something that is not yours or loose something during the event please
kindly contact the registration desk.
If you have any comments, questions or concerns (e.g. concerning radio/TV, religion and
culture, general rules, housing or conference events, …) you are welcome to write an email
to [email protected] or ask the local board at the conference venue.
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Insurance
All conference attendees are well advised to arrange private travel (including health!) insurance. The
conference organizers and committee accept no liability for personal accidents or damage to property while
in attendance at the conference and/or pre-conference workshop.
Banking
There are a lot of ATM machines and banks located around the University where you can withdraw money,
in particular found along Grindelallee.
Shopping
The shops in the city centre are generally open from 10 am- 8 pm. The supermarkets usually open between
7 am and 8 pm and there are some available that do not close before 10 pm. On Sundays the shops are
normally closed except for those at some stations.
Smoking
Smoking is prohibited by law in public areas/buildings within Germany. Ask whether your accommodation
bedroom is a smoking or non-smoking room. Please, make use of designated smoking areas outside public
buildings.
Communication
Only digital phones with GSM subscriptions and a roaming agreement will work in Germany. Visitors should
consult their supplier before departure. Common providers in Germany are Telekom, E-Plus, O2 and
Vodafone. Most, if not all, hotels offer WIFI access. All delegates will have free access to an Internet station
located in the conference center for the duration of the conference.
Currency
The Euro (€) is the single currency in Germany shared by most of the member states of the European Union,
which together make up the so-called “Eurozone”. Coin collectors may have a look at rare or uncommon
country-specific backsides of the Euro-coins. Credit cards such as VISA and MasterCard are widely accepted.
Traveller’s cheques are best exchanged at Hamburg Airport as many downtown banks do not exchange
them anymore.
Doctor and Pharmacy
“Ärztehaus an der Oper”, Dammtorstraße 27, 20354 Hamburg, close to the venue; www.aerztehaus-oper.de
There are a lot of pharmacies located around the University of Hamburg and the city centre.
Apotheke am Dermatologikum
Stephansplatz 1-5, 20354 Hamburg
Tel: +49 (0)40 81971960
Monday-Friday: 8:00-19:00
Saturday: 9:00-15:00
Enten Apotheke
Grindelallee 88-90, 20146 Hamburg
Tel: +49 (0)40 44140260
Monday-Friday: 8:00-19:00
Saturday: 9:00-14:00
Weather
Mild climate, the average temperature in September is 14 °C, i.e. 57 °F (low 9 °C, high 18 °C). Total rainfall in
September ± 70 mm, total rain days ± 11 out of 30.
Electric Current
220 Volts AV at 50 Hz. Do not forget to bring a proper universal non-grounded or grounded adapter plug for
use in Continental Europe.
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Emergency phone numbers
• Police: 110
• Fire Department: 112
• Ambulance: 112
Car hire at the airport
Avis
Budget
Enterprise Rent-A-Car
Europcar
Hertz
Sixt Mietwagen
Location: Terminal 2; contact: +49 (0)180 6557755, www.avis.de
Location: Terminal 2; contact: +49 (0)180 6217711, www.budget.de
Location: Terminal 2; contact: +49 (0)40 51316170, www.enterprise.de
Location: Terminal 2; contact: +49 (0)40 520 187 654, www.europcar.de
Location: Terminal 2; contact: +49 (0)40 59351367, www.hertz.de
Location: Terminal 2; contact: +49 (0)180 6252525, www.sixt.de
Bicycles
Computer-controlled stations for renting a bike, a so-called “StadtRad”, are available throughout the city,
including one next to the conference venue. You may rent a bike and connect it to an official station in the
city in the end of your tour. There are nice designated lanes for bicycles along most large streets in
Hamburg.
Parking
Parking is recommended at Congress Centre Hamburg (CCH) which offers 1150 parking spaces next to
Dammtor station for 2 €/hour or 14 €/day (open 24 hours).
There are also some possibilities for parking in the streets around the university main building but it is
probably difficult to get hold of a parking space there.
Taxi
There are many taxi services available in the city centre, at Dammtor station and also at the airport. If you
have any special requirements you can also call the following taxi services.
Taxi Hamburg: +49 (0)40 666666
Hansa Funktaxi: +49 (0)40 211211 or +49 (0)40 311311
Public Transport
Delegates should ensure when booking flights to Germany, they book to arrive in Hamburg Airport.
Hamburg City and Hamburg Airport are well serviced by public transport (both bus and rail).
For detailed information, please visit www.hvv.de.
Tipping
It is common to leave gratuity of about 8-10 % in restaurants and bars in Germany if you enjoyed your
food/drinks and the service.
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Hotel contact
name
Hotel NH Hamburg Horn
NH Hamburg Altona
NH Hamburg Mitte
NH Hamburg City
Mercure Hotel Hamburg City
Mercure Hotel Hamburg Mitte
Wyndham Garden Hamburg City Centre
Baseler Hof
Scandic Hamburg Emporio
email
[email protected]
[email protected]
[email protected]
[email protected]
[email protected]
[email protected]
[email protected]
[email protected]
phone
+49 (0)40 655970
+49 (0)40 4210600
+49 (0)40 441150
+49 (0)40 432320
+49 (0)40 236380
+49 (0)40 450691000
+49 (0)40 251640
+49 (0)40 359060
+49 (0)40 4321 870
Hotel am Rothenbaum
Hotel Engel Hamburg
Hotel Alster Hof
[email protected]
[email protected]
[email protected]
+49 (0)40 4153780
+49 (0)40 55 42 60
+49 (0)40 35 0070
Hotel Bellmoor
Hotel Hamburg-Alster Motel One
Hotel Ibis St.Pauli Messe
[email protected]
[email protected]
[email protected]
+49 (0)40 4133110
+49 (0)40 41924970
+49 (0)40 650460
Hotel Ibis Alster Centrum
[email protected]
+49 (0)40 248290
An overview of these recommended hotels is also available via www.iccbm15.org. Please ask the local board
if you need any help.
Shops at Hamburg central station are open seven days a week.
(photo as well as photos on pages 21, 28, 74, 160, 162 and 173 were taken
and modified by Sven Falke)
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Tourism information
With its 1.8 million inhabitants, Hamburg is a beautiful, maritime, modern, friendly and fascinating city. The
free and hanseatic city is located in the north of Germany, about 125 km from the North Sea. The world‘s
largest passenger aircraft, the A380, is manufactured in Hamburg and the city houses Europe's second
biggest harbor. Hamburg is famous for a wide range of arts and culture, city parks and beaches along the
rivers Elbe and Alster. In the historic center, the Town Hall, the churches and old ware- and storehouses, are
situated harmonically along with the HafenCity quarter. You may like to visit www.hamburg-travel.com or
http://english.hamburg.de/ for lots of additional information and suggestions.
Hamburg Card
The Hamburg Card not only offers you unlimited travel by bus, train and ferry, but also discounts for many
tours, museums, theaters, musicals, restaurants and all the major attractions in and around Hamburg.
The card is available for one day (9.50 €), three days (22.90 €) or five days (38.50 €).
For more information about the Hamburg Card, all attractions and tours and also for booking have a look
online.
Tours
• Hop on/hop off sightseeing tour
15 stops in a double-decker around the city center, the Alster, the Reeperbahn and the famous harbour. The
tour takes about 100 minutes, but you can also hop on and hop off at every station and continue the tour
afterwards. The buses depart every 30 minutes, tickets are available for 13 € per person and are valid for
the whole day. Audio commentary in different languages is available on request.
• XXL-harbour boat trip
Explore the variety of the port of Hamburg including the Altenwerder container terminal and the historic
warehouse complex on a 2 hour boat trip. Tickets cost approximately 22 €. Trips for 1 hour are available as
well.
• Alster boat trip on a historic steam boat
Experience the flair of Hamburg on Germany’s oldest steam boat during a 45 minute trip across the Alster.
The tour starts at the famous “Jungfernstieg” and then proceeds towards the outer Alster. Tickets are
available for 12.50 €.
Attractions
• St. Michaelis Church (Englische Planke 1, 20459 Hamburg)
The “Michel” is over 350 years old and Hamburg’s most famous church. Inside you will learn a lot about the
history of Hamburg. It is also possible to visit the crypt and ascent the viewing platform. You can either
choose the stairs or take the lift to the 106 metres high platform to enjoy a spectacular view over Hamburg.
A combined ticket for the tower and the crypt costs 6 €.
• Miniature Wonderland (Kehrwieder 2, 20457 Hamburg)
Discover the largest model railway system in the world in Hamburg’s famous Miniature Wonderland
museum (“Miniatur Wunderland”). Beside replicas of famous Hamburg districts and attractions, the
superlative model scenery includes for example landscapes of Austria, Scandinavia and the USA.
Tickets are available for 12 € per person.
• Hagenbeck Zoo (Lokstedter Grenzstraße 2, 22527 Hamburg)
The Hagenbeck Zoo accommodates more than 1,850 animals from all over the world. Additionally, you will
find an amazing variety of botanical species, protected views, open-air enclosures and numerous cultural
monuments. Discover the fascinating “Eismeer” where North Pole meets South Pole and also experience the
unique Tropical-aqaurium. Tickets: Zoological garden 20 €, tropical-aquarium 14 €, combo ticket 30 €.
Musicals
Hamburg is also famous for its variety of fantastic Musicals. The most popular and successful musicals
located in Hamburg are Disney’s The Lion King, the Phantom of the Opera and Rocky Hamburg.
For more information and tickets visit: www.hamburg-travel.com.
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Some recommended restaurants
Hofbräu an der Alster
Cuisine: Bavarian food
Location: Esplanade 6, 20354 Hamburg
Price category: €€*
Contact: +49 (0) 40 34993838
www.hamburg-hofbraeuhaus.de
Block House
Cuisine: Steakhouse
Locations: Jungfernstieg 1, 20095 Hamburg
Gänsemarkt Passage, 20354 Hamburg
Price category : €€*
Contact: +49 (0)40 30382215 (Jungfernstieg)
+49 (0)40 346005 (Gänsemarkt)
www.block-house.de
L’Osteria
Cuisine: Pizza and pasta
Location: Dammtorstr. 12, 20354 Hamburg
Contact: +49 (0)40 34106788
www.losteria.de
Bullerei
Cuisine: Variegated food
Location: Lagerstraße 34b, 20357 Hamburg
Price category : €€€*
Contact: +49 (0)40 33442110
www.bullerei.com
Blockbräu
Cuisine: Brewery pub
Location: Bei den St. Pauli-Landungsbrücken
3, 20359 Hamburg
Contact: +49 (0)40 44405000
www.block-braeu.de
Hard Rock Café Hamburg
Cuisine: American food
Location: Bei den St. Pauli-Landungsbrücken
5, 20359 Hamburg
Contact: +49 (0)40 4030068480
www.hardrock.com
Quán Dò Street Kitchen
Cuisine: Vietnamese food
Location: Georgsplatz 16, 20099 Hamburg
Price category : €*
Contact: +49 (0)40 32901737
www.quan-do.com
Mr. Cherng
Cuisine: Asian food and sushi
Location: Speersort 1, 20095 Hamburg
Contact: +49 (0)40 39870366
www.mr-cherng.de
Hans im Glück
Cuisine: Burgers and more
Location: Schäferkampsallee 1, 2035 Hamburg
Contact: +49 (0)40 41623667
www.hansimglueck-burgergrill.de
Mama trattoria
Cuisine: Italian food
Locations: Mittelweg 138, 20148 Hamburg
Schauenburgerstr. 44, 20095 Hamburg
Contact: +49 (0)40 53779799 (Mittelweg)
+49 (0)40 36099993 (Schauenburgerstr.)
www.mama.eu
SO NA MU
Cuisine: Korean food
Location: Grindelallee 89, 20146 Hamburg
Price category : €*
Contact: +49 (0)40 41308884
Many other restaurants providing for instance french, turkish or indian food or typical sea food are
around, close to the town centre, close to the university or around the harbour.
*the number of euro symbols (max. three) indicates the average price level
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Conference dinner (September 18th, 8 pm)
The Conference dinner will take place at the harbour on the museum ship “Rickmer
Rickmers” with a fantastic deck view over the port of Hamburg. A bus transfer is offered
for conference participants to go directly from the venue to the dinner location. Details
will follow on short notice. However, you may also like to choose another way or vehicle
to get there. If you go by public transport we suggest to exit the train (U-Bahn, yellow line
“U3“) at the station „Landungsbrücken“ directly at the harbour. As soon as you arrive at
the station you can already see the harbour and you are not far away from the large
“Rickmer Rickmers” which is predominantly green and red. You may cross the street
along this side of the harbour via a bridge and the destination is just a few hundred
metres away if you turn to the left (in the direction of the station „Baumwall“).
From the S-Bahn station „Dammtor“ that is next to the conference venue you may travel one
station to „Sternschanze“ (direction to „Altona“ or „Elbgaustraße“) and change the train by
entering U3 to go to „Landungsbrücken“ via „Feldstraße“ and „St. Pauli“. You may also travel
from “Dammtor” in the opposite direction to Central station („Hauptbahnhof“) and change
to the yellow line „U3“ there to get to the harbour and to „Baumwall“ and
„Landungsbrücken“.
Image © A. Conring
Location and contact:
Rickmer Rickmers Restaurant
At the St. Pauli Landungsbrücken - Ponton 1a / Fiete-Schmidt-Anleger
20359 Hamburg - Germany - tel. +49 (0)40 312 868;
http://www.rickmer-rickmers-gastronomie.de/rickmers_e/index.php
A map is available on the homepage.
19
DESY Campus map
20
(Notkestraße 85, 22607 Hamburg)
It is recommended to enter via the main gate close to
the workshop location and close to building 32c.
Main Gate
If you plan to visit DESY (Deutsches Elektronen-Synchrotron) and its unique radiation sources, the location
of the conference workshop, you may have a look at the map below to walk along the right way upon arrival
by bus or car. During the conference there will be a guided tour for a limited number of participants.
How to get access to the internet…
If you like to use the WLAN with your portable devices, please kindly register with your name at
the local registration desk for security reasons. Please follow the instructions below. Furthermore
you can receive an additional printed instruction how to connect and use your individual user
name and password.
As a requirement you have to customize the proxy server settings of your browser. Please select
the wireless network of the university with the SSID „GUEST“. Use the general WLAN password
„rzz-wlan“ to establish the connection. Upon starting your browser you will be directed to the
guest login page. You are asked to fill in your personal user name/password to finalize the
process.
Please logout at https://webauth.rzz.uni-hamburg.de/logout.html, press the “logout” button and
await the confirmation. The connection will stop automatically after 8 hours. You are asked to
restart the connection, if you like to continue.
Heinrich-Hertz-Turm, close to the venue, named after the German physicist Heinrich Hertz.
21
Detailed scientific conference program
17. 09. 2014
08:00 – 08:55
08:55 – 09:20
09:25 – 10:05
10:10 – 10:35
First day
Registration
Opening ceremony
ICCBM15 Organisers: C. Betzel & J.R. Mesters
Prof. C. Leopold (UHH vice president), Prof. E. Hartmann (UzL vice president),
Prof. E. Weckert (DESY board of directors)
Plenary/keynote: Leonid A. Sazanov
Crystallisation and structure of respiratory complex I, a giant molecular proton pump with
64 transmembrane helices and 9 Fe-S clusters
Coffee break
Membrane protein crystallization
Chair: L. DeLucas & H. Tidow
10:40 – 11:10 Vadim Cherezov
Microcrystal preparation for serial femtosecond crystallography in lipidic cubic phase
11:10 – 11:40 Poul Nissen
Lipid-based crystallization of membrane proteins
11:40 – 12:10 Hartmut Luecke
Membrane protein crystallization using cubic lipid phases, bicelles and vapor diffusion
12:10 – 12:30 Francoise Bonneté
Membrane protein crystallization in surfo: Influence of new surfactant structures on MP
phase diagrams
12:35 – 13:40
Lunch & exhibition
Crystallization for lead compounds in industry
Chair: M. Hennig & U. Egner
13:40 – 14:10 Jörg Benz
BACE2 crystal systems for drug discovery project support: Surface mutants and cocrystals
with Fab fragments, Fynomers and Xaperones
14:10 – 14:40 Linda Öster
Systematic studies on ligand solubility with the aim to increase the success rate of
obtaining protein-ligand complexes
14:40 – 15:10 Pamela Williams
Generating crystals for drug discovery programs: From initial chemical matter to candidate
15:15 – 15:45
Coffee & exhibition
Screening for crystals
Chair: J. Newman & S. Gournith
15:50 – 16:20 Fabrice Gorrec
Recent developments of initial crystallization screens for the determination of novel
protein
structures by X-ray diffraction
16:20 – 16:50 Simon Newstead
Membrane protein crystallisation and the development of MemGold, MemGold2 and the
MemAdvantage optimization screens.
16:50 – 17:20 Edward Snell
Comparing chemistry to outcome: A chemical distance metric, coupled with clustering and
hierarchal visualization applied to macromolecular crystallography
17:25 – 18:40
Poster Session & Exhibition
18:45
End of the first day
22
18. 09. 2014
08:15 – 08:55
Second day
Registration
Complementary methods
Chair: Th. Marlovits & A. Pearson
09:00 – 09:30 Trevor Forsyth
Integrated approached for the study of macromolecular systems
09:30 – 10:00 Anthony Watts
NMR crystallography of membrane proteins
10:00 – 10:25 Georgeos Skiniotis
Visualization of GPCR complexes by single particle EM
10:30 – 10:55
Coffee & exhibition
11:00 – 11:30 Jan-Pieter Abrahams
3D protein nano-crystallography by imaging and diffraction
11:30 – 12:00 Henning Stahlberg
High-throughput electron crystallography of 2D crystals of membrane proteins
12:00 – 12:55
Lunch, exhibition
Nucleation: Theory and practise
Chair: P. Vekilov & A. van Driessche
13:00 – 13:25 Dominique Maes
Nucleation precursors and the birth of crystals
13:25 – 13:50 Elias Vlieg
Growing high-resolution protein crystals using a simple, convection-free geometry
13:50 – 14:15 Jan Sedzik
Towards understanding nucleation and crystallization of proteins: Physico-chemistry of
crystallization drop
14:15 – 14:35 Dimitri Radajewski
Coupling droplet microfluidic and small angle X-ray scattering: Toward on-line
measurement of first nucleation step
14:40 – 15:45 Coffee, poster session & exhibition
In-vivo crystallization
Chair: M. Duszenko & L. Redecke
15:50– 16:15
Fasseli Coulibaly
Structural basis for the in vivo crystallization of viral proteins into functional assemblies
16:15 – 16:40 Cait MacPhee
What can bulk experiments tell us about nucleation in the confined environment of the
cell?
16:40 – 17:05 Daniel Passon
Organelles vs. whole cells approach: Serial femtosecond X-ray diffraction experiments on
in vivo grown peroxisomal methanol oxidase crystallites
17:10 – 17:25
Short break
Evening program
17:30 – 18:10 Keynote: Richard Giegé
20:00
Conference dinner
23
19. 09. 2014
08:15 – 08:55
Third day
Registration
Crystallization and scoring for SFX
Chair: H. Chapman & J. Helliwell
09:00 – 09:25 Andrew Aquila
Introduction to the European XFEL and its instrumentation for structural biology
09:25 – 09:50 Petra Fromme
Small is beautiful: How to grow nanocrystals for serial femtosecond crystallography
09:50 – 10:15 Guillermo Calero
Use of transmission electron microscopy in the discovery and optimization of protein nanocrystals
10:15 – 10:40 Gergely Katona
Picosecond dynamics of a photosynthetic reaction centre after photoactivation
10:45 – 11:15
Coffee, poster session & exhibition
Microfluidics crystallization, i.e. counter diffusion
Chair: J.A. Gavira & C. Sauter
11:20 – 11:45 Sachiko Takahashi
Efficient way to optimize protein crystallization condition for counter-diffusion method
11:45 – 12:10 Sarah Perry
Time resolved serial crystallography in a microfluidic device
12:10 – 12:35 Michael Heymann
Serial crystallography using a kinetically optimized microfluidic device for protein
crystallization and on-chip X-ray diffraction
12:40 – 13:35
Lunch, exhibition & poster session
Crystallization for neutron diffraction
Chair: D. Logan & W. Saenger
13:40 – 14:05 Zoë Fisher
Crystal growing and mounting techniques for macromolecular neutron crystallography: A
practical guide
14:05 – 14:30 Akihiko Nakamura
Crystallization phase diagram of glycoside hydrolase family 45 inverting cellulase for
neutron protein crystallography
14:30 – 14:55 Tobias Schrader
Requirements on crystal size at the new neutron diffractometer BioDiff
15:00 – 15:40
Coffee, poster session & exhibition
Microgravity
15:45 – 16:10
16:10 – 16:35
16:35 – 17:00
17:05 – 17:15
17:20 – 18:00
Chair: D.C. Yin & R. Giegé
Shigeru Sugiyama
Growth of protein crystals in hydrogels with high strength
Abel Moreno
Non-conventional techniques for protein crystallization and for separating nucleation and
crystal growth processes
Da-Chuan Yin
Improving protein crystal quality using diamagnetic levitation containerless method
Short Break
Keynote: Werner Kühlbrandt
What X-rays cannot do: Single-particle imaging, molecular tomography and the resolution
revolution in cryo-EM
24
20. 09. 2014
08:30 – 08:55
Fourth/last day
Registration office
Scoring and optimization in crystallization
Chair: R. Meijers & C. Wrenger
09:00 – 09:30 Janet Newman
Rules need tools / Tools need rules
09:30 – 10:00 Martin Caffrey
Crystallization of membrane proteins by the lipid cubic phase or in meso method. From
solubilized protein to final structure.
10:00 – 10:30 Bostjan Kobe
Linker-assisted crystallization of protein complexes: Application to TIR domains
10:35 – 11:05
Coffee break (optional poster removal)
11:10 – 11:40 Vasundara Srinivasan
Mitochondrial inner-membrane ABC transporter, ATM1 – crucial steps in crystallization and
structure determination
11:40 – 12:10 Alexandra Ros
Microfluidic tools for structure determination of membrane proteins
12:15 – 12:55
Keynote and closing lecture:
Christopher Tate
G protein-coupled receptors: Structure, function and drug development
13:00 – 13:15
Farewell address, end of conference
13:15
The end of ICCBM15
See you all in Prague for ICCBM16!
25
Exhibition
The exhibition within this conference will take place on the second (stand 1-5) and
first floor (stand 6-13) of the venue with companies from the scientific field taking
part as scheduled on the following two pages. A ground floor layout of the venue is
found on page 75.
no.
exhibitor
online
1
Bruker AXS GmbH
http://www.bruker.com/
2
Xtal Concepts
http://www.xtal-concepts.de/
3
Molecular Dimensions
http://www.moleculardimensions.com/
4
Douglas Instruments
http://www.douglas.co.uk/
5
TTP Labtech
http://ttplabtech.com/
26
no.
exhibitor
online
6
Formulatrix
http://www.formulatrix.com/
7
Labcyte
http://www.labcyte.com/
8
Wyatt
http://www.wyatt.eu/
9
Cube Biotech
http://www.cube-biotech.com/
10
Zinsser
http://www.zinsser-analytic.com/
11
Confocal Science
http://www.confsci.co.jp/
12
Dunn Labortechnik
http://www.dunnlab.de/
13
Rigaku
http://www.rigaku.com/
27
Oral presentations
City hall of Hamburg
28
Speakers (in alphabetical order)
name
institution
email
Universiteit Leiden, Netherlands
[email protected]
Prof.
Abrahams
Dr.
Aquila
Andrew
European XFEL GmbH, Germany
[email protected]
Dr.
Benz
Jörg
Roche AG, Swizerland
[email protected]
Dr.
Bonneté
Francoise
Université d'Avignon, France
[email protected]
Prof.
Caffrey
Martin
Trinity College Dublin, Ireland
[email protected]
Prof.
Calero
Guillermo
University of Pittsburgh, USA
[email protected]
Prof.
Cherezov
Vadim
The Scripps Research Institute, USA
[email protected]
Dr.
Coulibaly
Faselli
Monash University, Australia
[email protected]
Dr.
Fisher
Zoe
European Spallation Source, Sweden
[email protected]
Prof.
Forsyth
Trevor
Keele University, UK
[email protected]
Prof.
Fromme
Petra
Arizona State University, USA
[email protected]
Gorrec
Fabrice
MRC Laboratory of Mol. Biology, UK
[email protected]
Dr.
Heymann
Michael
CFEL, Germany
[email protected]
Dr.
Katona
Gergely
University of Gothenburg, Sweden
[email protected]
Prof.
Kobe
Bostjan
University of Queensland, Australia
[email protected]
Prof.
Kuehlbrandt
Werner
MPI für Biophysik, Germany
[email protected]
Prof.
Luecke
Hartmut
University of California, USA
[email protected]
Prof.
MacPhee
Cait
University of Edinburgh, UK
[email protected]
Dr.
Maes
Dominique
Vrije Universiteit Brussel, Belgium
[email protected]
Prof.
Giegé
Richard
University of Strasbourg, France
[email protected]
Prof.
Moreno
Abel
UNAM, Mexico
[email protected]
Jan-Pieter
Dr.
Nakamura
Akihiko
University of Tokyo, Japan
[email protected]
Dr.
Newman
Janet
CSIRO, Australia
[email protected]
Dr.
Newstead
Simon
University of Oxford, UK
[email protected]
Prof.
Nissen
Poul
aarhus university, Denmark
[email protected]
Dr.
Öster
Linda
AstraZeneca R&D Mölndal, Sweden
[email protected]
Dr.
Passon
Daniel
EMBL Hamburg, Germany
[email protected]
Dr.
Perry
Sarah
University of Chicago
[email protected]
Radajewski
Dimitri
LGC, France
[email protected]
Prof.
Ros
Alexandra
Arizona State University, USA
[email protected]
Prof.
Sazanov
Leonid
Medical Research Council, UK
[email protected]
Dr.
Schrader
Tobias
Forschungszentrum Jülich, Germany
[email protected]
Dr.
Sedzik
Jan
KTH, Sweden
[email protected]
Prof.
Skiniotis
Georgios
University of Michigan, USA
[email protected]
Dr.
Snell
Edward
Hauptman-Woodward Inst., USA
[email protected]
Dr.
Srinivasan
Vasundara
Loewe-Zentrum Synmicro, Germany
[email protected]
Prof.
Stahlberg
Henning
University of Basel, Switzerland
henning.stahlberg-at-unibas.ch
Prof.
Sugiyama
Shigeru
University of Tokushima, Japan
[email protected]
Dr.
Takahashi
Sachiko
Confocal Science Inc, Japan
[email protected]
Prof.
Tate
Christopher
MRC, UK
[email protected]
Prof.
Vlieg
Elias
Radboud University Nijmegen, NL
[email protected]
Prof.
Watts
Anthony
University of Oxford, UK
[email protected]
Dr.
Williams
Pamela
Astex Pharmaceuticals, UK
[email protected]
Prof.
Yin
Da-Chuan
N. Polytechnical University, China
[email protected]
29
Oral Presentation O-1
Abstract
Crystallisation and structure of respiratory complex I, a giant
molecular proton pump with 64 transmembrane helices and 9 Fe-S
clusters
Sazanov L
Mitochondrial Biology Unit, Medical Research Council, Cambridge, UK
[email protected]
NADH-ubiquinone oxidoreductase (complex I) is the first and largest enzyme in the
respiratory chain of mitochondria and many bacteria. It couples electron transfer between
NADH and ubiquinone to the translocation of four protons across the membrane. It is a
major contributor to the proton flux used for ATP generation in mitochondria, being one
of the key enzymes essential for life as we know it. Mutations in complex I lead to the most
common human genetic disorders. It is an L-shaped assembly formed by membrane and
hydrophilic arms. Mitochondrial complex I consists of 44 subunits, whilst the prokaryotic
enzyme is simpler and generally consists of 14 conserved “core” subunits of about 550
kDa in total. We use the bacterial enzyme as a “minimal” model to understand the
mechanism of complex I. We have determined all currently known atomic structures of
complex I, starting with the hydrophilic domain1,2, followed by the membrane domain3,4
and, finally, the recent structure of the entire Thermus thermophilus complex (536 kDa, 16
subunits, 9 Fe-S clusters, 64 TM helices)5, the largest asymmetric membrane protein
structure solved so far. Determination of the structure of the entire complex was possible
only through this step-by-step approach, building on from smaller subcomplexes towards
the entire assembly. As large membrane proteins are notoriously difficult to crystallize,
various non-standard and sometimes counterintuitive approaches6 were employed in
order to achieve crystal diffraction to high resolution and solve the structures. They
suggest a unique mechanism of coupling between electron transfer in the hydrophilic
domain and proton translocation in the membrane domain, via long-range (up to ~200 Å)
conformational changes. It resembles a steam engine, with coupling elements (akin to
coupling rods) linking parts of this molecular machine.
References
1
Hinchliffe P & Sazanov LA, Science 309, 771-774, (2005)
2
Sazanov LA & Hinchliffe P, Science 311, 1430-1436, (2006)
3
Efremov, RG, Baradaran R & Sazanov LA, Nature 465, 441-445, (2010)
4
Efremov, RG & Sazanov LA, Nature 476, 414-420 (2011)
5
Baradaran R et al., Nature 494, 443-448 (2013)
6
Sazanov LA et al., Biochem Soc Trans 41, 1265-1271 (2013)
30
Abstract
Microcrystal preparation for serial femtosecond crystallography in
lipidic cubic phase
Cherezov V
The Scripps Research Institute, USA
Recently we have established a new method of serial femtosecond crystallography in
lipidic cubic phase (LCP-SFX) for protein structure determination at X-ray free electron
lasers (XFEL). LCP-SFX uses gel-like lipidic cubic phase (LCP) as a matrix for growth and
delivery of membrane protein microcrystals to the intersection with an XFEL beam for
crystallographic data collection. In this talk I will describe the protocols for preparation
and characterization of microcrystals for LCP-SFX applications. The advantages of LCPSFX over traditional crystallography include the ability of high-resolution data collection
from sub-10 μm crystals of challenging membrane and soluble proteins at room
temperature with minimal effects of radiation damage, while eliminating the need for
crystal harvesting and cryo-cooling. Compared to SFX with microcrystals in solution using
liquid injectors, LCP-SFX reduces protein consumption by about two orders of magnitude
for data collection at 120 Hz.
31
Abstract
Lipid-based crystallization of membrane proteins
Nissen P
Aarhus University, Dept. Molecular Biology and Genetics
Centre for Membrane Pumps in Cells and Disease – PUMPkin, Danish National Research
Foundation
Danish Research Institute of Translational Neuroscience – DANDRITE, Nordic-EMBL
Partnership for Molecular Medicine
Membrane proteins are typically solubilized by detergents to facilitate purification and
crystallization. However, such procedures also delipidate the sample, and may lead to
inactivation of function and structures that are dominated by detergent interactions.
Traditionally, lipids were considered a source of inhomogeneity of membrane protein
samples, but over the recent years we have seen a progressive transition from detergentbased to lipid-based crystallization approaches. From our studies of ion pumps of the Ptype ATPase family we have observed critical involvement of lipid phases on function and
in crystallization. Numerous crystal forms have been obtained for a wide variety of P-type
ATPases and their different functional states. A recurring theme is a crystal packing based
on stacked bilayers, or so-called type I membrane protein crystals. These crystals forms
depend on a high content of solubilized lipids in the membrane protein samples, which
can either derive directly from solubilized, native membranes or from exogenous lipids
added to a purified and detergent-solubilized sample.
References
Nyblom M et al., Science 342, 123-7 (2013)
Gourdon P et al., Crystal Growth & Design 11, 2098–2106 (2011)
Quick M, Proc Natl Acad Sci U S A. 106, 5563-8 (2009)
Sorensen TL et al., J Biotechnol. 124, 704-16 (2006)
32
Abstract
Membrane protein crystallization using cubic lipid phases, bicelles
and vapor diffusion
Luecke H
Director, Center for Biomembrane Systems, University of California, Irvine.
The Cubic Lipid Phase (CLP) method for membrane protein crystallization has been refined to
allow large-scale screening of membrane proteins. Various parameters (CLP lipid, water content,
bilayer lipid additive, pH, ionic strength, precipitating agent etc.) can be varied. Several distinct
seven-transmembrane proteins were crystallized and their high-resolution structures determined.
In cases where the CLP method fails, the bicelle method or detergent-based methods were
employed to crystallize other membrane proteins.
Bacteriorhodopsin (BR): High-resolution maps from X-ray diffraction of bacteriorhodopsin crystal
obtained in CLP and some of its photointermediates have yielded insights to how the
isomerization of the bound retinal drives ion transport. Although some important mechanistic
details are still undecided, the events of the photochemical cycle are now understood to reflect
changes in specific hydrogen bonds of protein groups and bound water molecules in response to
motions of the retinal chain. A nearly complete lipid bilayer is also part of the x-ray-derived
atomic model. Surprisingly, we were unable to use the CLP method to obtain crystals of the
A215T mutant of BR, only the bicelle method provided reults.
Anabaena SR (ASR): The structure of a sensory rhodopsin from the cyanobacterium Anabaena has
been determined to 1.9 Å resolution using the CLP method. This represents the first eubacterial
rhodopsin structure. In comparison to the archaeal rhodopsins BR and SR there are many striking
rearrangements and shifts in hydrogen bonding patterns on both the extracellular and the
cytoplasmic half of the receptor. Also, the cytoplasmic face, which is thought to interact with the
soluble transducer, is structurally well defined and very different from that of the archaeal
rhodopsins. The structure of the soluble transducer of this photoreceptor (ASRT) has also been
determined - it forms a C4 tetramer with a new all-beta fold. Studies characterizing the interaction
between transmembrane photoreceptor and soluble transducer are underway.
Xanthorhodopsin (XR): a light-driven ion pump from the halophilic eubacterium Salinibacter ruber
found in saltern crystallizer ponds of España, contains a blue-absorbing carotenoid that functions
as a light-harvesting antenna for its retinal chromophore. This protein only crystallized from
bicelles. In addition to the adaptations to bind and accurately position the carotenoid antenna for
efficient excited-state energy transfer to the retinal, XR exhibits major structural differences to the
previously studied microbial pumps and photoreceptors. We also determined the ring structures
of a pentameric (C5) and a hexameric (C6) proteorhodopsin, from bicelles and CLP, respectively.
Lastly, I will present the results of structural & functional studies on a system responsible for acid
tolerance in certain bacteria. The system is able to maintain a periplasmic pH of ~6 even when the
medium has a pH of 2. The buffering system involves the enzyme urease that readily cleaves urea
into NH3 and CO2, which in turn act as a proton sink to reduce [H+] by four orders of magnitude.
This membrane protein crystallizes as a C6 hexamer using vapor diffusion after many rounds of
optimization.
33
Oral Presentation O-5 / P-105
Abstract
Membrane protein crystallization in surfo: Influence of new
surfactant structures on MP phase diagrams
Barret L-A1,2, Dahani M1, Raynal S1, Jungas C2, Polidori A1, Bonneté F1
1
2
Institut des Biomolécules Max Mousseron, Université d’Avignon, Avignon, France
Institut de Biologie Environnementale et de Biotechnologie, Saint-Paul-lez-Durance, France
Resolution of membrane protein (MP) structures at atomic level by crystallography remains a challenge of
structural biology because of the requirement of amphiphilic molecules (surfactant or lipids) in
crystallization processes, in surfo or in meso. However the use of hydrocarbonated surfactants such as DDM,
OG, in the different biochemical steps may destabilize MPs often leading to poor diffracting crystals. The
synthesis of new surfactants (1-3) with more rigid or bulkier hydrophobic chain (cyclic or hemifluorinated)
could be an interesting alternative to usual detergents to grow diffracting crystals.
New surfactants used for stabilization and crystallization of MPs: PCC-α−Malt (1), F6diGluM (2) and F2H9-β−malt (3)
Here we present a study about the influence of hydrophobic chain structure on crystallization of a
photosynthetic complex, RC-LH1-pufX from Rhodobacter blasticus. Micelle assemblies were characterized by
small angle X-ray scattering (SAXS) and form factor model fitting (SASfit). Crystallization was approached
by weak interaction characterization between surfactant micelles, via second virial coefficient
measurements using SAXS or DLS. This approach allowed us to correlate surfactant phase diagrams and
crystallization of RC-LH1-pufX (4).
Variation of second virial coefficient of PCCαM vs DDβM as a function of PEG; Crystals of RC-LH1 in the two surfactants;
Residual bound lipids in the two surfactants
MP structural stability and therefore quality diffraction depending on residual MP-bound lipids, nature and
amount of residual lipids were characterized using high performance thin layer chromatography (HPTLC)
for RC-LH1 purified in DDM and PCCM (5). We found more residual lipids bound to the protein when
prepared in the most hydrophobic surfactant.
In conclusion, these new surfactants appear to have a great potential for stabilization and crystallization of
membrane proteins in surfo.
References
1.
2.
3.
4.
5.
Hovers J et al., Molecular Membrane Biology, 28(3), 171-81 (2011)
Abla M et al., J. Fluor. Chem 134, 63-71 (2012)
Bonneté F et al., in preparation
Barret L-A et al., J. Phys. Chem. B, 117(29), 8770-81 (2013)
Barret L-A et al., J Chromatography A, 1281, 135-141 (2013)
34
Abstract
BACE2 crystal systems for drug discovery project support: Surface
mutants and cocrystals with Fab fragments, Fynomers and
Xaperones
Benz J
Roche Pharma Research and Early Development, Small Molecule Research, Molecular
Design and Chemical Biology, Roche Innovation Center Basel, 4070 Basel, Switzerland
[email protected]
Iterative cocrystal structures of target proteins in complex with ligands form the basis for
rational drug design. However, obtaining cocrystal structures is often difficult or
impossible for certain ligands, even when crystallization of the protein has already been
well established. Robust crystallization systems are thus required that reproducibly and
rapidly yield well diffracting crystals of all desired protein–ligand complexes. The aspartic
protease BACE2 is responsible for the shedding of the transmembrane protein Tmem27
from the surface of pancreatic -cells, which leads to inactivation of the -cell proliferating
activity of Tmem27. This role of BACE2 in the control of -cell maintenance suggests
BACE2 as a drug target for diabetes. High-resolution BACE2 ligand cocrystal structures
would contribute significantly to the investigation of this enzyme as either a drug target or
anti-target. Surface mutagenesis, BACE2-binding antibody Fab fragments, single-domain
camelid antibody VHH fragments (Xaperones) and Fyn-kinase-derived SH3 domains
(Fynomers) were used as crystallization helpers to obtain the first high-resolution
structures of BACE2. The different strategies used for raising and selecting BACE2 binders
for cocrystallization will be described and the crystallization success, crystal quality and
the time and resources needed to obtain suitable crystals will be compared.
Reference
Banner DW et al., Acta Cryst. D69, 1124-37 (2013)
35
Abstract
Systematic studies on ligand solubility with the aim to increase the
success rate of obtaining protein-ligand complexes
Öster L1, Jonasson E1, Eriksson P-O1, Grönberg G2, Sigfridsson K3, Sandmark J1
1Structure
2CVMD
& Biophysics, Discovery Sciences, AstraZeneca R&D Mölndal, Sweden
iMed, AstraZeneca R&D Mölndal, Sweden
3Pharmaceutical
Development, AstraZeneca R&D Mölndal, Sweden
Structure determination of ligands bound to their target proteins using X-ray crystallography is the core of
structure based drug discovery. However, complex structures are not always easy to achieve, especially in
cases where the affinity is poor and the solubility of the ligand is low. The solubility of the ligand in the
soaking solution is critical for success and can be manipulated in different ways, for example by including
different cosolvents, additives and varying soaking temperatures. However, the lack of systematic studies
makes it hard to device efficient strategies.
The aim of this study was to systematically test the effect of different cosolvents, vehicles and soaking
temperatures on the solubility of the ligands in the soaking condition. To investigate this we have developed a
method to measure the concentration of the ligands in the soaking experiment using Nuclear Magnetic
Resonance (NMR). We compared the results to the ability to obtain protein-ligand structures using X-ray
crystallography in a system where the success rate to obtain protein-ligand complex structures has been
limiting.
The different cosolvents tested were water, DMSO and a mixture of DMA/PEG400/H20. We also dissolved the
ligands using cyclodextrin and nanosuspensions - two common approaches for in vivo drug administration.
Furthermore, we tested dissolving the ligands by addition of either HCl or NaOH depending on the ligand
properties. The soaking temperatures investigated were 20 and 30°C respectively. We have further explored
the physicochemical properties of the ligands and correlated those values to our results with the aim to
facilitate prediction on which solubilisation method to use for each individual ligand.
The results from this study will be presented together with further discussion on how we work with ligands
with low solubility to obtain protein-ligand structures.
References
Hassell A, et al., Acta Crystallographica section D 63, 72-79 (2007)
Sigfridsson K, et al., Asian journal of Pharmaceutical Sciences 3, 161-168 (2009)
Sigfridsson K, et al., European Journal of Pharmaceutics and Biopharmaceutics 67, 540-547 (2007)
36
Abstract
Generating crystals for drug discovery programs: From initial
chemical matter to candidate
Williams PA1
1Astex
Pharmaceuticals, Cambridge CB4 0QA, UK
The use of protein structure data can vary widely in drug discovery projects, with different
organisations using the technique at different stages in a campaign. At Astex
Pharmaceuticals access to crystal structures is fully integrated into, and a fundamental
part of, the drug discovery process. Crystal structures are used to identify initial chemical
hits as part of a biophysical screening campaign and in the later stage optimisation of
protein-ligand interactions. Generation of crystals for use in fragment screening presents
different challenges to those encountered in novel structure determination, challenges
which include reproducing large numbers of crystals with consistent diffraction in the
presence of high concentration ligands.
In this talk I will discuss the properties you might look for in a crystallographic system
used in the drug discovery process, how to achieve these and illustrate how this varies
from target to target.
37
Abstract
Recent developments of initial crystallization screens for the
determination of novel protein structures by X-ray diffraction
Gorrec F1
1MRC
Laboratory of Molecular Biology, Cambridge, England
Since protein samples are increasingly challenging to produce, a common approach to initial crystallization screening is
to employ a limited number of trials (96-192 conditions). In addition, since no predictions can be made about
crystallization, conditions are usually selected empirically [Jancarik & Kim, 1991]. The corresponding minimal screens
are regularly optimized following approaches biased towards the type of samples investigated, statistics and costs
[Kimber et al., 2003; Rupp & Wang, 2004; Newman et al., 2005]. Subsequently, there are now a plethora of screens that
have been made commercially available (each screen typically includes 96 conditions).
Increasing the initial yield of quality crystals has a multitude of positive impacts on the overall process of structure
determination by X-ray diffraction, for example, if diffraction quality is sufficient, it reduces the need for subsequent
optimization. Also, as reproducibility may be an issue, it is advantageous to collect as much data as possible during the
initial screening.
At the MRC-LMB, despite using a set of commercially available conditions considered as very large ≥1440 conditions)
[Stock et al., 2005], the yield of crystals with sufficient quality to solve structures is generally very low (<1%). In
addition to sample properties, an underlying reason for such low yield is the very large number of combinations of
variables associated with macromolecular crystallization [McPherson, 1990].
We argue that our approach to initial screening is still under-sampled [Gorrec, 2013] and hence an even larger set of
conditions should be preferred whenever possible. As a consequence, we are constantly investigating novel
formulations to expand the scope of our initial screening such as the MORPHEUS and Pi screens [Gorrec, 2009; Gorrec et
al., 2011]. Instead of selecting conditions empirically, screens are designed de novo in order to avoid bias and attempt
crystallization with increasingly challenging novel samples.
The formulation of a Pi screen is derived from the incomplete factorial approach [Carter & Carter, 1979]. Combinations
of 3 reagents are produced following the modular distribution of a given set of up to 36 stock solutions. Maximally
diverse conditions can be produced by taking into account the properties and the concentrations of the chemicals used
to formulate a 96-condition screen.
MORPHEUS, also consisting of 96-conditions, integrates several innovative approaches developed elsewhere, such as
mixes of potential ligands [McPherson & Cudney, 2006], convenient buffer systems [Newman, 2004] and precipitant
mixes that also act as cryoprotectants. Molecules frequently observed in the Protein Data Bank (PDB) to co-crystallize
with proteins have been selected to formulate MORPHEUS.
The MORPHEUS and Pi screens improved our yield of quality crystals. In addition, after commercialization of the Pi and
MORPHEUS screens, it was observed that a multitude of other laboratories have also benefited from their formulations.
These results give further grounds to our initial claim that screens currently available are under-sampled. It also shows
the importance of employing innovative approaches to further developing new screens.
References
Carter CW Jr & Carter CW, J. Biol. Chem. 254, 12219-12223 (1979)
Gorrec F, J. Appl. Cryst. 42, 1035-1042 (2009)
Gorrec F et al., Acta Cryst. D67, 463-470 (2011)
Gorrec F, J. Appl. Cryst. 46, 795-797 (2013)
Jancarik J & Kim SH, Acta Cryst. D24, 409-411 (1991)
Kimber MS et al., Proteins 51, 562-586 (2003)
McPherson A, Eur. J. Biochem. 189, 1-23 (1990)
McPherson A & Cudney B, J. Struct. Biol. 156, 387-406 (2006)
Newman J, Acta Cryst. D60, 610-614 (2004)
Newman J et al., Acta Cryst. D61, 1426-1431 (2005)
Rupp B & Wang J, Methods, 390-407 (2004)
Stock D et al., Prog. Biophys. Mol. Biol. 88, 311-327 (2005)
38
Abstract
Membrane protein crystallisation and the development of
MemGold, MemGold2 and the MemAdvantage optimization screens
Newstead S
Department of Biochemistry, University of Oxford, Oxford, OX1 3QU, UK.
In this talk I will highlight the latest trends in alpha helical membrane protein
crystallization using information gathered from the Protein Data Bank. Every month we
update our in house database, which contains the relevant information on crystallization
experiments from recently published studies. We have used this resource to develop a
number of targeted sparse matrix style crystallization screens, including MemGold &
MemGold2, in addition to a recently developed optimization screen MemAdvantage. The
strategy behind the design of these screens and their optimal use will be discussed.
Information is also gathered on heavy atom phasing for alpha helical MPs, and I will
present our latest analysis on successful strategies recently reported in the literature.
39
Abstract
Comparing chemistry to outcome: A chemical distance metric,
coupled with clustering and hierarchal visualization applied to
macromolecular crystallography
Bruno AE1, Ruby AM1, Luft JR2,3, Grant TD2, Seetharaman J4, Hunt JF4,
Montelione GT5,6, Snell EH2,3
1Center
for Computational Research, State University of New York (SUNY), Buffalo, NY 14260, USA;
Medical Research Institute, Buffalo NY 14203, USA; 3SUNY Buffalo Dept. of
Structural Biology, Buffalo NY 14203, USA; 4Department of Biological Sciences, The Northeast Structural
Genomics Consortium, Columbia University, New York, NY 10032, USA; 5Northeast Structural Genomics
Consortium, Department of Molecular Biology and Biochemistry, Center for Advanced Biotechnology and
Medicine, Rutgers, The State University of New Jersey, Piscataway, NJ 08854, USA; 6Department of
Biochemistry, Robert Wood Johnson Medical School, University of Medicine and Dentistry of New Jersey,
Piscataway, NJ 088546, USA.
2Hauptman-Woodward
Many bioscience fields employ high-throughput methods to screen multiple biochemical conditions. The
analysis of these becomes tedious without a degree of automation. Crystallization, a rate limiting step in
biological X-ray crystallography, is one of these fields. Screening of multiple potential crystallization
conditions (cocktails) is the most effective method of probing a proteins phase diagram and guiding
crystallization but the interpretation of results can be consuming. To aid this empirical approach a cocktail
distance coefficient was developed to quantitatively compare macromolecule crystallization conditions and
outcome. These coefficients were evaluated against an existing similarity metric developed for
crystallization, the C6 metric, using both virtual crystallization screens and by comparison of two related
1,536-cocktail high-throughput crystallization screens. Hierarchical clustering was employed to visualize
one of these screens and the crystallization results from an exopolyphosphatase-related protein from
Bacteroides fragilis, (BfR192) overlaid on this clustering. This demonstrated a strong correlation between
certain chemically related clusters and crystal lead conditions. While this analysis was not used to guide the
initial crystallization optimization, it led to the re-evaluation of unexplained peaks in the electron density
map of the protein and to the insertion and correct placement of a sodium, potassium and phosphate atoms
in the structure. With these in place, the resulting structure of the putative active site demonstrated features
consistent with active sites of other phosphatases which are involved in binding the phosphoryl moieties of
nucleotide triphosphates. The new distance coefficient, CDcoeff, appears to be robust in this application and
coupled with hierarchical clustering and the overlay of crystallization outcome reveals information of
biological relevance. While tested with a single example the potential applications related to crystallography
appear promising and the distance coefficient, clustering and hierarchal visualization of results undoubtedly
have applications in wider fields.
(i)
(ii)
(iii)
Figure 1. (i) Heatmaps illustrating the results of the Hierarchial Clustering Algorithm on 1,536 different
chemical cocktails used for crystallization screening, (ii) isolation of one cluster from screening of BfR192
illustrating the crystallization results and (iii) the structure of the BfR192 exopolyphosphatase-related
protein showing the two domains and highlighting a cleft containing sodium, potassium and four phosphate
ions identifed through re-analysis following clustering demostrating that crystallization screening can also
supply functional information.
40
Abstract
Integrated approached for the study of macromolecular systems
Forsyth T
Biochemistry Department, South Parks Road, Oxford, OX1 3QU, UK
41
Abstract
NMR crystallography of membrane proteins
Watts A
Biochemistry Department, South Parks Road, Oxford, OX1 3QU, UK
Solid state NMR methods are very versatile with respect to sample geometry and form1,
and both amorphous and (2D and 3D1) crystalline materials can be studied to give high
resolution spectra of use in deriving structural models. Being a short-range technique
(dipolar interactions exist over Ås only), these crystals can be very small (nano- or
amorphous), and certainly much smaller than can be manipulated for conventional single
crystal diffraction approaches – powder samples still contain short-range order. Indeed,
biomolecules that crystallize into sub-µm crystals but of little use for diffraction, can be
ideal for study by solid state NMR. Here some examples will be given of how we have
devised methods and resolved high-resolution structural information from both 2D and
3D crystals of membrane proteins2,3,4. Sensitivity enhancements using dynamic nuclear
polarization (DNP), are also possible with crystalline sample form5. Both back- bone2, and
ligand/prosthetic group details6,7 have been obtained, usually ahead of the XRD or EM
structures – some comparison will be presented where the crystal structures have been
subsequently resolved.
References
1.
2.
3.
4.
5.
6.
7.
Watts et al., Methods in Molecular Biology – Techniques in Protein NMR Vol. 278 (ed.
K. Downing), Humana Press, New Jersey, pp. 403-474 (2004)
Higman, et al., Angew. Chem. Int. Ed. 50, 8432-5 (2011)
Varga et al., Biochim. Biophys. Acta 1768, 3029-3035 (2007)
Varga et al., J. Biomol. NMR 41, 1-4 (2008)
Pike et al, J. Mag. Res. (front cover) 215, 1-9 (2012)
Ulrich et al., Nature Structural Biology 2, 190-192 (1995)
Watts, Nature Reviews Drug Discovery 4, 555-568 (2005)
42
Abstract
Visualization of GPCR complexes by single particle EM
Skiniotis G
Life Science Institute, University of Michigan, USA
Single particle electron microscopy (EM), devoid of the need for large-scale sample
preparations or protein crystallization, has been established as a very powerful approach
for the 3D structural characterization of biological macromolecular complexes. Recent
advances in instrumentation and image reconstruction algorithms have not only enabled
high resolution structure determination by this methodology, but also the analysis of
conformational dynamics within the same particle population, thereby providing crucial
insights to mechanistic aspects of protein function. While discussing the basics of this
application, we will describe single particle EM visualization on GPCRs and their
complexes, as exemplified in a GPCR/G protein complex, a GPCR/arrestin complex, and a
class C GPCR. We will further discuss the hybridization of EM data with other biophysical
and biochemical methods, as well as the current challenges and future directions.
43
Abstract
3D protein nano-crystallography by imaging and diffraction
Abrahams JP, Nederlof I, Clabbers M, Li Y-W, Genderen E
Leiden Inst. of Chemistry, Universiteit Leiden, Einsteinweg 55, Leiden, The Netherlands
Many proteins do not form crystals large enough for 3D structure determination by X-ray
crystallography. For sub-micron sized 3D protein crystals there are several alternatives:
intense XFEL sources, electron diffraction and electron imaging. Recently, known crystal
structures of 3D sub-micron protein crystals could be refined using XFEL and electron
diffraction data. Solving new structures using these approaches remains a problem, as abinitio crystal phasing is hampered by experimental constraints that severely compromise
data quality. These constraints include incomplete sampling of Bragg spots and (for
electron diffraction), dynamic and inelastic scattering.
In the case of electron diffraction, we show that complete sampling of Bragg spacings can
be achieved with a fast TimePix quantum area detector in combination with continuous
crystal rotation. This detector is of unprecedented sensitivity. We could collect up to 30
Degrees of data from a single frozen lysozyme crystal of only 100x200x200 nm3.
Furthermore, we demonstrated that in the case of pharmaceuticals, we could collect a
complete data set at room temperature from crystals of only 20x20x20 nm2 to 0.8 Å
resolution, and solve the structure.
We also show that ab-initio phase information can be measured directly by electron
imaging of nano-crystals. We have shown that this experimental approach also allows
enhancing the resolution of electron-images of 3D nano-crystals to beyond 2 Angstrom
including crystal phases, by separating mosaic blocks within nano-crystals through image
processing.
44
Abstract
High-throughput electron crystallography of 2D crystals of
membrane proteins
Stahlberg H
Center for Cellular Imaging and NanoAnalytics (C-CINA), Biozentrum, University of Basel,
Switzerland
Email: [email protected]
Electron Crystallography uses cryo-transmission electron microscopy (cryo-EM) to study
the structure of membrane proteins that are lipid membrane reconstituted and twodimensionally crystallized.
This lecture will present structural studies by cryo-EM of MloK1, a cyclic-nucleotide
modulated potassium channel that features ligand binding domains and also putative
voltage sensor domains. The structure of the lipid-membrane reconstituted bacterial
channel was determined in the presence and absence of its ligand cAMP, revealing
significant movements of the ligand binding domains, which interact with the channel’s
putative “voltage sensor domains”, thereby presumably altering the channel at the
selectivity filter domain [1].
This lecture will also discuss recent method developments in streamlined data collection
using a direct electron detector camera, and image processing advances, including a single
particle approach to 2D crystal images, as well as movie-mode image processing for 2D
crystal data.
References
Kowal J et al., Nature Communications 5, 3106 (2014)
45
Abstract
Nucleation precursors and the birth of crystals
Maes D1, Sleutel M1, Van Driessche AES1, Giglio M2, Potenza M2, Sanvito T2, Stiens J3,
Zhang Y3, Vorontsova MA4, Vekilov PG4.
1Structural
Biology Brussels, Vrije Universiteit Brussel, Brussels, Belgium; 2Physics Department,
University of Milano, Milan, Italy; 3Department of Electronics and Informatics, Vrije Universiteit
Brussel, Brussels, Belgium; 4Department of Chemical and Biomolecular Engineering and
Department of Chemistry, University of Houston, Houston, USA
Nucleation of crystals is one of the most secretive processes in chemistry, materials
science, and biophysics. The two-step mechanism of nucleation was first put forward for
protein crystallization and subsequently its applicability was demonstrated for colloids,
small-molecule organic and inorganic substances, and polymers. The nucleation
precursors are liquid-like protein rich clusters with a size that varies from under one
hundred to several hundred nanometers. The mesoscopic clusters exist with similar
characteristics both in the homogeneous region of the phase diagram of the protein
solution (where no condensed phases, liquid or solid, are stable or present as long-lived
metastable domains) and under conditions supersaturated with respect to the ordered
solid phase.
In this lecture we will first focus on the characteristics of the nucleation precursors and
their role in the nucleation process. Experiments show that the presence of clusters has a
large effect on crystal nucleation. Clusters exhibit an Ostwald ripening behavior during a
crystallization cycle. With Brownian Microscopy, previously used to study the dynamics of
the clusters as a whole, we obtained information on the dynamics within the clusters.
Furthermore our experiments revealed that the assimilation of clusters by the growing
crystals can trigger a self-purification cascade of impurity-poisoned crystal surfaces.
Finally we will report on our endeavors to capture crystals at birth with more novel
techniques such as ‘polarized Brownian Microscopy’, Confocal Depolarized Dynamic Light
Scattering and TeraHertz spectroscopy.
46
Abstract
Growing high-resolution protein crystals using a simple,
convection-free geometry
Vlieg E1, Adawy A1, Törnroth-Horsefield S2, de Grip W2, van Enckevort W1
1Radboud
University Nijmegen, The Netherlands, 2University of Gothenburg, Sweden
The growth of high quality single protein crystals, yielding the highest X-ray resolution,
remains a bottleneck in macromolecular crystallography. Convection-free conditions
should lead to improved crystal quality, because in such a case growth is slow and the
formation of defects and the incorporation of impurities minimized. Convection-free
conditions have been achieved in the microgravity environment of space and in magnetic
fields, but the impact on protein crystal growth has been limited due to the experimental
difficulties with these methods.
Here we show that through a laboratory-based setup (Adawy, 2013), an entirely
convection-free crystallization environment is achieved. This is accomplished by using an
upside-down geometry, where crystals grow at the ‘ceiling’ of a growth cell completely
filled with the crystallization solution. The ‘ceiling crystals’ experience the same diffusionlimited conditions as in space microgravity environments. The new method was tested on
bovine insuline and two hen egg-white lysozyme polymorphs. In all cases, ceiling crystals
diffracted X-rays to resolution limits beyond their current world records, even while
solutions of moderate purity were used.
We further used the ceiling geometry for a direct comparison of the impurity
incorporation between growth with and without convection under otherwise identical
conditions. Several impurities were tested, and in most cases diffusion-limited growth
leads to crystals with lower impurity levels and/or improved diffraction quality, even for
heterogeneous impurities.
References
Adawy et al., Cryst. Growth Des., 13, 775 (2013)
47
Abstract
Towards understanding nucleation and crystallization of proteins:
Physico-chemistry of crystallization drop
Sedzik J
Royal Institute of Technology, Department of Chemical Engineering and Technology,
Protein Crystallization Facility, Stockholm, Sweden and Department of Applied Chemistry,
School of Engineering
The University of Tokyo, Tokyo, Japan. [email protected],
[email protected],
Physico-chemistry has its special terminology which has to be defined at the beginning of
each presentation. The terms: aggregation, nucleation or crystallization are intuitive and
self-explanatory. There is more complex vocabulary: like “surface tension”, “interfacial
tension”, “interfacial energy”, “contact angle”, “entropy” or “free Gibbs energy”. Some of
them are more difficult to understand and will be accurately defined during presentation
The basic equation for formation of single nucleus followed by crystallization is described
as
Where
G(volume) – change in free
2
energy per unit volume (erg/cm ), r – radius of formed nucleus (in cm), σ – surface
tension (dyna/cm) or surface energy per unit area or interfacial energy (erg/cm2).
ΔG total – total energy needed when forming nucleus of diameter r and further growth of
crystal (erg).
From this fundamental equality the following equation can be derived:
r’ – critical radius for nuclei
G’– energy barrier formation of nucleus
These fundamental equations cover homo- and heterogeneous nucleation. In this
presentation the model of crystallization experiments is considered which resulted in
crystals or precipitate. The calculated interfacial tension between crystallization medium
and surface of protein crystal is calculated allowing for better insight on nucleation and
crystallization. The measurement of contact angle (mimics interaction of mother liquor
with surface of protein crystal) allowing calculating of the energy barrier and critical
radius for formation of the nucleus.
Acknowledgment
This work was supported by the Japanese Society of Promotion of Science (Tokyo, Japan)
as a BRIDGE Fellowship (RC 2092910) and short term JSPS Invitation Fellowship
Programs for Research in Japan.
48
Abstract
Coupling droplet microfluidic and small angle X-ray scattering:
toward on-line measurement of first nucleation step
Radajewski D1, Pham NV1, Bonneté F2, Guillet P2, Brennich ME3, Round A4,5, Pernot
P3, Biscans B1, Teychené S1
1Laboratoire
de Génie Chimique UMR 5503, Université de Toulouse; Toulouse, France
des Biomolécules Max-Mousseron, Université dʼAvignon, Avignon, France
3 European Synchrotron Radiation Facility, Grenoble, France
4 European Molecular Biology Laboratory, Grenoble Outstation, France
5 Unit for Virus Host-Cell Interactions, Univ. Grenoble Alpes-EMBL-CNRS, France.
2 Institut
Small Angle X-Ray Scattering (SAXS) is a powerful technique to characterize structure and form factors of
colloidal systems in solution. The combination of microfluidics and SAXS provides a powerful tool to
investigate kinetics and transitions at different molecular levels and relevant timescales. During the past
decade, given the growing interest of the microfluidic community to use in-situ characterization techniques,
significant progresses have been made for coupling microfluidics and X-Ray scattering. However, very few
studies deal with the combination of SAXS and digital microfluidic. In digital microfluidic the sample is
compartmentalized inside droplets. Each droplet acts as a microreactor in which the operating conditions
(concentration, pH, and temperature) can be finely tuned, without cross-contamination. Beyond the
opportunity of manipulating sub-microliter of fluids, the major interest in coupling SAXS and droplet
microfluidics for protein crystallization is to continuously flow thousands of microdroplets of sample
through the X-Ray beam for exploring phase diagrams from undersaturated to supersaturated conditions.
This present study focused on the development of a robust digital microfluidic experimental set-up to probe
protein interactions in solution for nucleation studies. Microfluidic chips were designed to adapt to the
sample holder on the BioSAXS beamline BM29 at ESRF (Grenoble) (Left figure); a fluorinated surfactant was
especially synthesized and diluted in a fluorinated carrier oil, together resisting to radiation damage. Protein
droplets were therefore generated in a capillary by mixing stock solution of protein, buffer and precipitant
in the carrier oil and then exposed to X-ray beam. Two proteins have been tested, lysozyme and rasburicase.
We clearly show, first, that the scattering curves of two proteins obtained in nanoliter droplets fit well with
their reference scattering curves (using CRYSOL) proving the stability of proteins in fluorinated droplet and
to radiation damage (Fig in the middle). In addition, by continuously changing the relative flow rate of the
different stock solutions in the microfluidic chip, we could measure in-situ lysozyme interactions in solution
(Right figure) and can now consider protein nucleation study.
30
Rasburicase (1r51.pdb)
Lysozyme with salt
25
I(q)/c (a.u.)
I(q)/c (a.u.)
100
10
1
0 mM NaCl
200mM NaCl
20
500mM NaCl
750mM NaCl
15
10
5
0.1
0
1
q (nm-1)
2
3
0
0
1
q (nm-1)
2
3
Left: Experimental microfluidic set up on BM29; Middle: Measured and calculated scattering curves of rasburicase by SAXS
in microfluidic droplets; Right: Scattering curves of Lysozyme in different salt concentrations showing attractive
interactions.
49
Abstract
Structural basis for the in vivo crystallization of viral proteins into
functional assemblies
Boudes M1, Aizel K1, Garriga D1, Hijnen M1, Olieric V2, Schulze-Briese C2, Bergoin M3,
Coulibaly F1
1School
of Biomedical Sciences, Monash University, Clayton, Australia
Light Source at Paul Scherrer Institut, Villigen, Switzerland
3Université Montpellier 2, Montpellier, France
2Swiss
Despite significant theoretical and experimental advances, protein crystallization remains
the main bottleneck in most structure determination pipelines. Over the last decade, the
study of proteins that crystallize as part of the normal viral infectious cycle has provided
new insights into this process that may ultimately contribute to alleviating this bottleneck.
The first X-ray structures of in vivo crystals were elucidated from viral polyhedra produced
by cypoviruses [1] and baculoviruses [2] using microcrystals directly purified from
infected insects. The function of polyhedra is to package up to hundreds of viral particles
constituting the main infectious form of the virus and allowing the virus to persist for
years in the environment like bacterial spores.
By contrast with these viruses, insect poxviruses produce two types of microcrystals in
infected cells: virus-containing spheroids representing their main infectious form; and
spindles, bipyramidal crystals of the viral fusolin protein, that contribute to the oral
virulence of these viruses. We have recently determined the structures of these functional
crystals allowing a comparative analysis of the four completely different architectures of in
vivo crystals. These various molecular organisations suggest common strategies of selfassembly that hint to features facilitating crystallization and more particularly in vivo
crystallization, which has been recently proposed as a potential new route for structure
determination [3].
1.
2.
3.
Coulibaly F et al., Nature 446, 97–101 (2007)
Coulibaly F et al., Proc Natl Acad Sci 106, 22205–10 (2009)
Koopmann R et al., Nat Methods 9, 259–62 (2012)
50
Abstract
What can bulk experiments tell us about nucleation in the confined
environment of the cell?
MacPhee C
University of Edinburgh, UK
In small volumes, the kinetics of protein self-assembly is expected to show significant
variability, arising from intrinsic molecular noise. We have recently developed a simple
stochastic model including nucleation and autocatalytic growth which allows us to predict
the effects of molecular noise on the kinetics of autocatalytic self-assembly. Our analysis
shows that significant lag-time variability can arise from both primary nucleation and
from autocatalytic growth and provides a way to extract mechanistic information on earlystage aggregation from bulk experiments. Within the confined environment of the cell, the
kinetics of self-assembly are influenced by the locally crowded environment. I will
describe how this will influence the kinetics and outcome of self-assembly.
51
Abstract
Organelles vs. whole cells approach:
Serial femtosecond X-ray diffraction experiments on in vivo grown
peroxisomal methanol oxidase crystallites
Passon DM1, Jakobi AJ1,2, Knoops K4, Seine T1, Liang M3, Stellato F3, White TA3,
Komadina D1, Messerschmidt M5, Bourenkov G1, van der Klei I4, Chapman HN3,
Wilmanns M1
1
European Molecular Biology Laboratory, Hamburg Unit, Notkestrasse 85, 22603 Hamburg, Germany. 2
European Molecular Biology Laboratory, Meyerhofstr. 1, 69117 Heidelberg, Germany. 3 Center for FreeElectron Laser Science (CFEL), Deutsches Elektronensynchrotron (DESY), Notkestrasse 85, 22607 Hamburg,
Germany. 4 Molecular Cell Biology, Groningen Biomolecular Sciences and Biotechnology Institute (GBB),
University of Groningen, Groningen, The Netherlands. 5 SLAC National Accelerator Laboratory, 2575 Sand
Hill Road, Menlo Park, California 94025, USA
Peroxisomes are membrane-enclosed organelles of eukaryotic cells harboring more than fifty reactions,
including lipid metabolism and the scavenging of reactive oxygen species. Peroxisomes are capable of
containing an unusually high load of proteins, which can result in the in situ crystallization of these proteins,
as observed in several yeast species, plants and vertebrates (1-6). In the yeast Hansenula polymorpha for
example alcohol/methanol oxidase (AO) is the predominant peroxisomal protein and it oligomerises into a
670 kDa complex that spontaneously forms nano-sized crystallites within peroxisomes under specific
growth conditions (1-3).
For a proof-of-principle demonstration of peroxisomal in situ crystallization, we used the H.
polymorpha AO. The lattice of naturally derived AO crystals within peroxisomes has been characterized (23). The apoprotein form of AO crystals diffracts to maximally 6 Å resolution at synchrotron-based X-ray
sources. A high resolution X-ray structure has not yet been determined. Owing to the limited size of H.
polymorpha peroxisomes (less than 1 µm) and the formation of in situ crystals, serial femtosecond X-ray
crystallographic experiments at X-ray free electron lasers will be ideally suited for solving the structures of
these crystals.
For serial femtosecond X-ray diffraction experiments at the CXI end station of LCLS we
sampled the whole peroxisome organelle as well as the whole H. polymorpha cells. Prior to the experiments,
the samples were pre-characterized and the buffer conditions were adapted for liquid jet applications. Peakfinding routines mining the collected scattering profiles of the whole cell suspensions identified ~4500
Bragg-sampled diffraction patterns. Virtual powder patterns assembled from the individual frames match
low-resolution powder diffraction patterns of concentrated suspensions of purified peroxisomes collected
with maximum photon flux at the P14 beamline at the PETRAIII synchrotron, confirm that the observed FEL
diffraction patters result from Bragg diffraction of peroxisomal crystallites. Currently, the maximum
resolution achieved in the diffraction patterns is limited to 20-15 Å. Future work will need to address
improved sample preparation protocols in order to assess whether diffraction to a resolution sufficient to
permit structure solution can be obtained.
References
1. Veenhuis M et al., Mol. Cell. Biol. 1, 949–957 (1981)
2. Vonck J and van Bruggen EF, J. Bacteriol. 174, 5391–5399 (1992)
3. van der Klei IJ, et al., Yeast 7, 15–24 (1991)
4. Frederick SE & Newcomb EH, J Cell Biol. 43(2), 343-353 (1969)
5. Heinze M et al., Cryst Res Technol. 35(6-7), 877-886 (2000)
6. Zaar K, Voelkl A, Fahimi HD, J Cell Biol. 113(1), 113-121 (1991)
52
Abstract
Giegé R
University of Strasbourg, France
To be announced.
53
Abstract
Introduction to the European XFEL and its instrumentation for
structural biology
Aquila A
European XFEL GmbH, Hamburg, Germany
The under-construction European XFEL will become the highest brilliance X-ray source in
the world when it comes online in just a few short years. As has already been
demonstrated [ref-Boutet, et al], XFELs offer a new world of opportunity for structural
biology applications, potentially overcoming the need for large crystals (or perhaps even
crystals at all), and avoiding much of the issues of radiation damage. The unprecedented
repetition rate of the XFEL.EU provides benefits that other XFEL facilities cannot presently
match.
This presentation will provide an update of the construction status of the European XFEL,
its basic parameters and a (very brief) overview of the planned instrumentation for
structural biology.
54
Abstract
Small is beautiful: How to grow nanocrystals for serial femtosecond
crystallography
Fromme P et al (see refs for full list of authors)
Arizona State University, Department of Chemistry and Biochemistry, Tempe, USA
The method of Serial Femtosecond nanocrystallography (SFX) provides a novel concept for
structure determination, where X-ray diffraction “snapshots” are collected from a fully hydrated
stream of protein nanocrystals, using femtosecond pulses at a high energy X-ray free-electron
laser, the Linac Coherent Light Source [1],[2],[3]. In SFX diffraction data of nanocrystals are
collected at room temperature in liquid jet. As femtosecond pulses are briefer than the time-scale
of most damage processes, femtosecond nanocrystallography overcomes the problem of X-ray
damage in crystallography [4]. SFX also provides also the unique perspective to for time resolved
femtosecond nanocrystallography [5,6] towards molecular movies of light-driven reactions. This
talk will feature different methods that can be used to grow and characterize nanocrystals of high
quality and uniform size distribution for SFX studies [7],[8]. The talk will focus on the question
how one can identify nanocrystals in high throughput in broad crystallization screens and
describe methods how to scale up production of nanocrystals for Serial Femtosecond
Crystallography. The talk will also emphasize what conditions have to be met for successful crystal
delivery and data collection on the nanocrystal samples at Free Electron Lasers.
References:
[1] Chapman HN et al. Nature 470, 73-77 (2011)
[2] Redecke L et al., Nature 339, 227-230 (2013)
[3] Liu W et al. Science 342, 1521-1524
[4] Barty A et al, Nature Photonics 6, 35–40 (2012)
[5] Aquila A et al., Optics Express. 20(3), 2706-16 (2012)
[6] Kupitz C et al., Nature, July 9 (2014)
[7] Hunter MS, Fromme P, Methods 55, 387-404 (2011)
[8] Kupitz C et al., Philos Trans R Soc Lond B Biol Sci 369, 20130316 (2014)
Acknowledgement: Experiments were carried out at the Linac Coherent Light Source and the
Advanced Light Source, operated by Stanford University and the University of California
(respectively) on behalf of the U.S. Department of Energy (DOE), Office of Basic Energy Sciences.
We acknowledge support from DOE through the PULSE Institute at the SLAC National Accelerator
Laboratory and by Lawrence Livermore National Laboratory under Contract DE-AC5207NA27344, the National Institute of Health (awards 1R01GM095583-01 and NIGMS PSI:Biology
grant U54 GM094599); the NSF BioXFEL Science and Technology center MCB1021557, the
National Science Foundation (awards 0417142 and 1021557) and the the NSF BioXFEL Science
and Technology center MCB1021557 , the Center for Bio-Inspired Solar Fuel Production, an
Energy Frontier Research Center funded by the U.S. Department of Energy, Office of Science, Office
of Basic Energy Sciences under Award Number DE-SC0001016 and the European Research
Council (AXSIS grant).
55
Abstract
Use of transmission electron microscopy in the discovery and
optimization of protein nano-crystals
Stevenson H1, Lin G1, Barnes C1, Kovaleva E2, DePonte D2, Reynolds S1, Calero M1, Fu
X1, Makhov A1, Soltis M2, Cohen A*2 and Calero G*1
1Department
of Structural Biology, University of Pittsburgh School of Medicine. 2Stanford
Synchrotron Radiation Laboratory, Stanford University
X-ray Crystallography experiments remain the most successful method to obtain
structural information from biological targets, comprising approximately 80% of the
current protein data base entries. Advances in molecular biology and biochemical
techniques have allowed expression and purification of a significant number of new
protein targets including human and pharmacological relevant proteins. Similarly,
innovations in synchrotron science including development of micro-focused x-ray beams
and advances in detector technologies as well as development of user-friendly
crystallography software packages have expedited crystal-to-structure time frames.
However, non-withstanding the abundance of targets or the increasingly faster crystal-tostructure pipeline, crystallization of protein targets remains the most significant
bottleneck in structure determination by X-ray crystallography.
Recent developments at the Linac-Coherent Light Source (LCLS) at Stanford University
have demonstrated that it is now feasible to solve crystal structures using diffraction data
from low-micrometer (<15 microns) or nanometer sized crystals. These groundbreaking
innovations open the door to obtain atomic information from those protein samples
whose intrinsic disorder prevented formation of large visible crystals. Using transmission
electron microscopy (TEM), we show that: 1) protein NCs are readily detected in most
crystallization samples; 2) TEM allows quantitative evaluation of NC lattices to determine
their potential diffraction quality; and 3) TEM can be used as tool to guide/improve
crystallization. Given the new opportunities that X-FEL offers to the field of
crystallography, and that NCs are ubiquitous, efficient methodologies to detect high quality
(well diffracting) NCs will be essential for its future development.
56
Abstract
Picosecond dynamics of a photosynthetic reaction centre after
photoactivation
Arnlund D1, Johansson LC1, Wickstrand C1, Barty A2, Williams GJ3, Malmerberg E1, Davidsson J4,
Milathianaki D3, DePonte DP2,3, Shoeman RL5,6, Wang D7, James D7, Katona G1, Westenhoff S1 , White
TA2, Aquila A2, Bari S6,8, Berntsen P1, Bogan M9, Brandt van Driel T10 , Doak RB5,7, Kjær KS10,11, Frank
M12, Fromme R13, Grotjohann I13, Henning R14, Hunter MS13 , Kirian RA7, Kosheleva I14, Kupitz C13,
Liang M2, Martin AV2, Nielsen MM10, Messerschmidt M2,3, Seibert MM3, Sjöhamn J1, Stellato F2,
Weierstall U7, Zatsepin NA7 , Spence JCH7 , Fromme P13 , Schlichting I5,6, Boutet S3, Groenhof G15,16,
Chapman HN2,17,18, Neutze R1
1 Department of Chemistry and Molecular Biology, University of Gothenburg, Gothenburg, Sweden.
2 Center for Free-Electron Laser Science, Deutsches Elektronen-Synchrotron DESY, Hamburg, Germany.
3 Linac Coherent Light Source, Stanford Linear Accelerator Center (SLAC) National Accelerator Laboratory,
Menlo Park, CA, USA.
4 Department of Photochemistry and Molecular Science, Uppsala University, Uppsala, Sweden
5 Max-Planck-Institut für medizinische Forschung, Heidelberg, Germany.
6 Max Planck Advanced Study Group, Center for Free Electron Laser Science, Hamburg, Germany.
7 Department of Physics, Arizona State University, Tempe, Arizona, USA.
8 Max-Planck-Institut für Kernphysik, Heidelberg, Germany.
9 Photon Ultrafast Laser Science and Engineering Center Institute, Stanford Linear Accelerator Center
(SLAC) National Accelerator Laboratory, Menlo Park, California, USA.
10 Department of Physics, Technical University of Denmark, Lyngby, Denmark.
11 Niels Bohr Institute, University of Copenhagen, Copenhagen, Denmark.
12 Lawrence Livermore National Laboratory, Livermore, California, USA.
13 Department of Chemistry and Biochemistry, Arizona State University, Tempe, Arizona, USA.
14 BioCARS, The University of Chicago, Illinois, USA.
15 Nanoscience Center, University of Jyväskylä, Jyväskylä, Finland.
16 Department of Chemistry, University of Jyväskylä, Jyväskylä, Finland.
17 Department of Physics, University of Hamburg, Hamburg, Germany.
18 Centre for Ultrafast Imaging, Hamburg, Germany.
The active state of proteins is often very far from thermal equilibrium. One remarkable example is the
photosynthetic reaction centre of purple bacteria, which non-destructively thermalize the excess energy of
visible photon. This process is crucial for maintaining its structural integrity as the excess energy is
sufficient to unfold the protein. Here we present a detailed structural and kinetic model of transient energy
transfer following excitation using time-resolved wide angle X-ray scattering at an X-ray free electron laser.
The data can be described by a minimal set of four components: an ultrafast component (C1) (peaks at 1 ps),
a protein component (C2) (7 ps), which displays oscillatory features characteristic of protein difference
scattering, and a non-equilibrated (C3) (14 ps) and equilibrated (C4) heating components which reaches
maximum amplitude in about 100 ps. Notably, the protein structural changes precede the heating
components providing a unique insight to the well ordered process of thermalization in biological systems.
Basis spectra of the spectral decomposition derived from
time dependent difference wide-angle X-ray scattering
signal.
References
Arnlund D, et al., Nat Methods doi: 10.1038/nmeth.3067 (2014)
57
Abstract
Efficient way to optimize protein crystallization condition for
counter-diffusion method
Takahashi S
Confocal Science Inc., Tokyo, Japan
Counter-diffusion (CD) method is a simple and useful technique for protein
crystallization (Zeppezauer et al., 1968; Salemme, 1972; García-Ruiz and
Moreno, 1994; Otálora et al., 2009). One of the major CD methods is called
“gel-acupuncture method”; a protein solution is loaded into a capillary,
which is stuck into a gel layer, and a reservoir solution is poured on the gel
layer. Both the solutions diffuse out and in the capillary through the gel, so
the concentration gradients of the solutions are formed in the capillary
from the opposite direction.
We developed another type of CD method, “gel-tube method” (Tanaka et al.,
2004). We use a piece of gel-tubing instead of the gel-layer. We have been
using it for more than ten years and have applied it to more than 500
protein samples. This method inherits all the merits of the gel-acupuncture
method, and, in addition, it is easy to prepare and is useful for post-soaking
of ligands and/or for cryoprotectant.
Two advantages of CD method compared to a vapour-diffusion (VD)
method are introduced here.
The first advantage is that a gradual supersaturation gradient is formed along the capillary as diffusion proceeds, so a
wide range of crystallization conditions can be explored in one single experiment. And the concentration change in the
capillary is milder than the VD method, so crystals can grow in the lower supersaturation regions, resulting in growing
fewer numbers and less clustered crystals of better diffraction quality. One-dimensional (1-D) simulation of the
diffusion profiles of the protein and the reservoir solutions along the capillary can help optimize the crystallization
condition (Tanaka et al., 2004). Also to know the solution condition around the crystal by 1-D simulation is
indispensable for harvesting and cryocooling crystals for avoiding crystal deterioration. However, because of the slow
diffusion, sometimes crystals don’t grow in CD method even though they grow in VD method. To solve this problem, the
protein solution and the reservoir solution are mixed and gel tube is pre-soaked in the reservoir solution beforehand.
These help rise the precipitant concentration in the capillary much faster, resulting in the increase of the crystal growth
probability. If it does not work effectively, seeding technique can be adopted. Micro-seeds are prepared using small
crashed crystals of the target protein and are loaded after the protein solution in the capillary. As a result, the crystal
growth is observed almost certainly in CD method.
The other advantage of CD method is that the solution in the capillary is not concentrated but is gradually replaced by
the reservoir solution, so that the concentration and the composition of the solution in the capillary can be controlled.
Therefore, no phase separation occurs in the capillary which often occurs in VD especially when membrane protein is
solved in the detergent solution. It is because the detergent is too much concentrated in the VD drop. However, this
feature of CD method sometimes evokes some trouble if we apply the protein and the reservoir solutions for VD method
to CD method. We noticed that the protein sample sometimes contains significant amount of the salt which is essential
for the crystallization, but its concentration in the capillary was not enough increased as that in the drop of VD method.
We would like to emphasize that the optimum amount of salt as well as significant amount of dominant precipitant
might be essential for protein crystal growth (Collins, 1997; 2004; Inaka et al., 2008). Rough estimation method of the
optimum salt concentration in the reservoir solution for CD method, which may help saving time and cost for obtaining
fine crystals, will be introduced (Yamanaka et al., 2011).
This work was supported by Japan Aerospace Exploration Agency High-quality Protein Crystal Growth Experiment
(JAXA PCG).
References
Collins, KD, Biophys. J., 72, 65-76 (1997)
Collins, KD, Methods, 34, 300-311 (2004)
García-Ruiz, JM, Moreno, A, Acta Cryst., D50, 484–490 (1994)
Inaka, K, et al., Acta Cryst., A64, C584 (2008)
Otálora, F, et al., Prog. Biophys. Mol. Biol., 101, 26–37 (2009)
Salemme, FR, Arch. Biochem. Biophys., 151, 533 (1972)
Tanaka, H, et al., J. Synchrotron Rad., 11, 45–48 (2004)
Yamanaka, R, et al., J. Synchrotron Rad., 18, 84–87 (2011)
Zeppezauer, M, et al., Arch. Biochem. Biophys., 126, 564-573 (1968)
58
Abstract
Time resolved serial crystallography in a microfluidic device
Perry SL1,2,, Srajer V3,4, Pawate AS2, Guha S2, Schieferstein J2, Kenis PJA2
1Department
of Chemical Engineering, University of Massachusetts Amherst, Amherst,
MA, USA;
of Molecular Engineering, University of Chicago, Chicago, IL, USA;
3Consortium for Advanced Radiation Sources, University of Chicago, Chicago, IL, USA;
4Department of Chemical & Biomolecular Engineering, University of Illinois, Urbana, IL,
USA; 5Renz Research Inc., Westmont, IL, USA
2Institute
Renewed interest in room temperature diffraction has been prompted by the desire to
observe structural dynamics of proteins as they function. Serial crystallography, an
experimental strategy that aggregates small pieces of data from a large, uniform pool of
crystals, has been demonstrated at synchrotrons and X-ray free electron lasers. We utilize
a microfluidic crystallization platform for serial Laue diffraction from macroscopic
crystals. This strategy takes advantage of integrated microfluidics and the exquisite
control over transport phenomena possible at the microscale to enable the growth of a
large number of high quality, isomorphous crystals for serial diffraction. The ability to
perform in situ diffraction directly in our microfluidic devices also eliminates the need for
manipulation or harvesting of the fragile protein crystals, while serial data collection
avoids challenges associated with radiation damage by enabling the collection of as little
as a single frame of diffraction data from an individual crystal.
Here we report the in situ time-resolved structural analysis of photoactive yellow protein
(PYP), performed entirely in a microfluidic chip. Following laser-initiation of the protein
photocycle, we observed the evolution of structural changes over timescales ranging from
nanoseconds to milliseconds. A complete dataset was obtained by merging small slices of
Laue data taken from many individual crystals. Electron density difference maps
generated from merged data highlight the expected conformational changes of the
chromophore with time, validating our approach.
Looking forward, the ability of our microfluidic platforms to grow and collect in situ
crystallographic data from a large number of protein crystals has the potential to enable
the use of synchrotron-based Laue diffraction for both serial crystallography, and the
dynamic structural study of biochemical reactions that are functionally irreversible due to
factors such as X-ray radiation damage, slow reversion back to the ground state,
limitations of the crystal lattice, or the actual irreversible nature of the reaction.
Ultimately, the most significant potential of our microfluidics-based approach comes from
the integration of fluid handling capabilities to enable flow-cell based triggering of
enzymatic reactions for dynamic crystallographic analysis.
59
Abstract
Serial crystallography using a kinetically optimized microfluidic
device for protein crystallization and on-chip X-ray diffraction
Heymann M1,2,#,*, Opthalage A2,*, Wierman JL3, Akella S2,‡, Gruner SM3,4,5,6, Fraden S2
∗contributed
equally to this work
Program in Biophysics and Structural Biology, Brandeis University, Waltham, MA, USA; 2Martin
Fisher School of Physics, Brandeis University, Waltham, MA, USA, #present address: Center for FreeElectron Laser Science, DESY, Hamburg, Germany; 3Field of Biophysics, Cornell University, Ithaca, NY, USA;
‡present address: Collective Interactions Unit, Okinawa Institute of Science and Technology Onna-Son,
Okinawa, Japan; 4Cornell High Energy Synchrotron Source (CHESS) and Macromolecular Diffraction Facility
at CHESS (MacCHESS), Cornell University, Ithaca, NY, USA; 5Department of Physics, Cornell University,
Ithaca, NY USA; 6Kavli Institute at Cornell for Nanoscale Science, Cornell University, Ithaca, NY, USA
1Graduate
We developed an emulsion based serial crystallographic technology in which nanoliter sized droplets of
protein solution are encapsulated in oil and stabilized by surfactant. Once the first crystal in a drop is
nucleated, the small volume generates a negative feedback mechanism that lowers the supersaturation,
which we exploit to produce one crystal per drop. We diffract, one crystal at a time, from a series of room
temperature crystals stored on an X-ray semi-transparent microfluidic chip and obtain a complete data set
by merging single diffraction frames taken from different unoriented crystals. As proof-of-concept we solved
the structure of glucose isomerase to 2 angstroms resolution, demonstrating the feasibility of highthroughput serial X-ray crystallography using synchrotron radiation.
(A) We used a frame to hold the X-ray transparent chip. (B) X-ray chip mounted on the goniometer inside
the Cornell CHESS F1 beamline. (C) Glucose isomerase crystals inside of the microfluidic device. Using a
motorized stage, each crystal can be centered in the collimated X-ray beam. The beam is 100 µm in
diameter. (D) Representative diffraction pattern of a glucose isomerase crystal taken at room temperature
from inside the chip. Crystals diffracted to 1.4 angstrom resolution with a mosaicity as low as 0.04o. The
bottom right quadrant shows the diffraction pattern after background subtraction.
60
Abstract
Crystal growing and mounting techniques for macromolecular
neutron crystallography: A practical guide.
Fisher SZ1, E. Oksanen E1, A. Hiess A1.
1Scientific Activities Division, Science Directorate, European Spallation Source, P.O. Box
176, Lund 22100, Sweden.
To date there are only ~65 neutron crystal structures deposited in the Protein Data Bank.
This is in stark contrast to the remaining >100,000 structures that are mostly from X-ray
crystallography, NMR spectroscopy, EM etc. About 40% of these neutron structures were
determined from samples prepared at the Protein Crystallography Station (PCS) support
labs at Los Alamos National Laboratory. The main deterrent for most researchers to
pursue a neutron structure of their favourite target is sample size and lack of
instrumentation. There are two new instruments that have come on-line recently:
IMAGINE and MaNDi at Oak Ridge National Laboratory. So while more instruments are
steadily being added to the global pool, sample size is still the biggest bottleneck for
successful protein neutron crystallography experiments. The average required protein
crystal volume is still in the ~1 mm3 range on most reactor and spallation neutron
sources. There are a handful of examples of successful structure determinations from
samples smaller than 0.3 mm3, but these are the exception. Large crystal growth is difficult
to achieve for multiple reasons. It is challenging and expensive in practice to produce
several 100 mgs of pure protein for neutron experiments. Conditions that work well for
routine crystal growth (for X-ray applications) may be have to adapted and then rescreened in scaled-up, large drops. Of course this consumes large amounts of precious
sample. If large, single crystals can be grown, these have to be mounted and H/D
exchanged in quartz capillaries at least 3-4 weeks prior to beam time. This step typically
happens well in advance of awarded or anticipated beam time and requires that samples
are quite stable, possibly for several months. Despite the availability of more neutron
instruments, these are heavily oversubscribed and users can wait quite a long to get beam
time. Due to this, samples often have to be prepared will in advance of a scheduled
experiment. Large crystal growth, mounting, and the in-capillary H/D exchange takes a
long time, thus a successful neutron experiments requires careful planning and sample
management. This presentation will go over some practical approaches and methods that
have been used successfully at the PCS and support labs for user several projects. This will
include details of drop and crystal scale up, crystal feeding, materials required for large
crystal growth and mounting as well as details about the PCS instrument and sample
requirements.
61
Abstract
Crystallization phase diagram of glycoside hydrolase family 45
inverting cellulase for neutron protein crystallography
Nakamura A1, Ishida T1, Kusaka K2, Tanaka I2, Yamada T2, Fushinobu S1, Niimura N2, Igarashi K1, and
Samejima M1
1Graduate School of Agricultural and Life Sciences, The University of Tokyo
2Frontier Research Center for Applied Atomic Sciences, Ibaraki University
Neutron protein crystallography (NPC) is a powerful tool for determining the hydrogen position and water
orientation in proteins (Niimura N & Podjarny A, 2011), but a much larger crystal of protein is needed for
NPC than for X-ray crystallography because the brightness of the neutron beam is weak. Thus crystal
preparation is a major bottleneck of NPC.
Endoglucanase (EC 3.2.1.4) belonging to glycoside hydrolase (GH) family 45 in the carbohydrate-active
enzymes database (Henrissat B. et al., 1991) is an enzyme produced by various organisms. This family has
been divided into three sub-families (A, B and C) from the phylogenetic analysis (Igarashi et al., 2008),
whereas the detailed character has been demonstrated only for the sub-family A cellulase from Humicola
insolens (HiCel45A). This enzyme has a pair of carboxylic residues located in the substrate-binding cleft, and
it hydrolyzes cellulose via an inverting mechanism (Davies et al., 1995), in which one of the carboxylic
residues acts as a general base and the other as a general acid. We recently discovered that Cel45A from the
basidiomycete Phanerochaete chrysosporium (PcCel45A, sub-family C) has only a glutamic acid residue at
the appropriate position for the general acid and no other acidic residue around catalytic center, but it has
activity (Igarashi et al., 2008). Therefore, we considered that NPC analysis of PcCel45A would give new
insights to the reaction mechanism of this enzyme.
To grow large protein crystals, it is necessary to know the properties of the target protein in the
crystallization solution. Here, a crystal preparation method of PcCel45A is reported, guided by the
crystallization-phase diagram. Nucleation and precipitation conditions were determined by sitting-drop
vapor diffusion. Saturation and undersaturation conditions were evaluated by monitoring crystal
dissolution, and a crystallization phase diagram was obtained. To obtain a large crystal, crystallization
solution was prepared on a sitting bridge (diameter = 5 mm). Initial crystallization conditions were 40 ml of
crystallization solution (40 mg/ml protein with 30.5% 3-methyl-1,5-pentane-diol in 50 mM tris-HCl pH 8.0)
with a 1000 µl reservoir (61% 3-methyl-1,5,- pentanediol in 50 mM tris-HCl pH 8.0) at 293 K. After the first
crystal appeared, the concentration of precipitant in the reservoir solution was reduced to 60% to prevent
formation of further crystals. Finally, we obtained a crystal of 6 mm3 in volume (3 mm x 2 mm x 1 mm),
which was suitable for neutron diffraction.
The crystallization phase diagram of PcCel45A and grown crystal: The crystallization phase diagram
(protein concentration vs precipitant concentration) was determined at 293 K in 50 mM tris-HCl buffer pH
8.0. Triangle: unsaturated; circle: saturated; diamond: nucleation; square: precipitation.
References
Niimura N & Podjarny A, Neutron Protein Crystallography Hydrogen, Protons, and Hydration in Bio-macromolecules,
Oxford University Press (2011)
Henrissat B. et al., Biochem. J. 280, 309-316 (1991)
Igarashi, K. et al., Appl. Environ. Microbiol. 74, 5628–5634 (2008)
Davies, GJ et al., Biochemistry, 34, 16210–16220 (1995)
62
Abstract
Requirements on crystal size at the new neutron diffractometer
BioDiff
Schrader TE1, Ostermann A2, Monkenbusch M1,3, Laatsch B4, Jüttner P2, Petry W2 and
Richter D1,3
1In
Jülich Centre for Neutron Science JCNS, Forschungszentrum Jülich GmbH, Outstation at MLZ, Garching,
Germany; 2Heinz Maier-Leibnitz Zentrum (MLZ), Technische Universität München, Garching, Germany;
3Institute for Complex Systems ICS, Forschungszentrum Jülich GmbH, Jülich, Germany;
4Forschungszentrum Jülich GmbH, Zentralinstitut für Engineering, Elektronik und Analytik, Engineering und
Technologie (ZEA-1), Jülich, Germany
Protein crystallography using neutrons as a probe has two major advantages as compared
to employing x-rays: First the hydrogen atoms can be seen even at moderate resolutions of
2.5 Å. Therefore, protonation states of amino acid side chains can be determined. Also, the
exact orientation and hydrogen bond pattern of well ordered water molecules can be
inferred. The hydrogen bonding of ligands to the active centre of the protein can also be
investigated. The second important advantage of neutrons as compared to x-rays is that
photo-reduction of metallo-proteins does not occur to a considerable amount and the
metallic centre can be observed in its native oxidation state. One major draw back of
neutron protein crystallography is the need for fairly large crystals usually of more than
one mm3 in Volume. It is sometimes challenging to obtain crystals of that size. When
intermediate states shall be cryo-trapped rapid freezing of these crystals imposes another
challenge. In this contribution some considerations are presented on how to obtain mm3
crystal sizes.
To reduce incoherent scattering background stemming mostly from hydrogen atoms it is
necessary to replace all exchangeable hydrogen atoms with deuterium. This is achieved by
carefully exchanging the mother liquor with a deuterated one. Suitable procedures for
achieving that will be discussed. In some cases it may even be advisable to use
perdeuterated proteins expressed in deuterated minimal media.
With respect to a potential measurement at the instrument BioDiff, hints are given on how
to prepare your crystals at the home lab for a room temperature or a cryo-crystallography
experiment. Depending on the unit cell size I discuss the requirements on the size and the
need for deuteration of the protein crystal. BIODIFF is designed as a monochromatic
instrument with a wavelength spread of less than 3 %. Its main advantage is the possibility
to adapt the central wavelength to the size of the unit cell of the sample crystal while
operating with a clean monochromatic beam that keeps the background level low.
Time permitting I will shortly introduce other neutron crystallography beamlines e.g.
LADI-III and D16 at ILL, MaNDi at SNS, IMAGINE at HIFR and iBIX in Japan. Also, a new
Korean instrument has been taken into operation recently. The protein crystallography
station (PCS) at LANCSE and the planned ESS instrument for protein crystallography will
be reported on in another contribution.
63
Abstract
Growth of protein crystals in hydrogels with high strength
Sugiyama S1,8, Matsumura H2,5, Sazaki G2,3, Maruyama M2, Takahashi Y2,
Yoshikawa H4, Yoshimura M2, Adachi H2,5, Takano K5,6, Murakami S5,7, Inoue T2,5,
Mori Y2,5
1Graduate
School of Science, Osaka University, Osaka, Japan; 2Graduate School of Engineering, Osaka
University, Osaka, Japan; 3The Institute of Low Temperature Science, Hokkaido University, Sapporo, Japan;
4Department of Chemistry, Saitama University, Saitama, Japan; 5SOSHO, Inc., Osaka, Japan; 6Graduate School
of Life and Environmental Sciences, Kyoto Prefectural University, Kyoto, Japan; 7Graduate School of
Bioscience and Biotechnology, Tokyo Institute of Technology, Yokohama, Japan; 8JST, ERATO, Lipid Active
Structure Project, Toyonaka, Japan
Crystal structure analysis of biological macromolecules has an essential role in the development of new
pharmaceutical compounds by structure-based drug design and the study of various vital phenomenon.
After successful crystallization experiments, protein crystals must be captured the crystals on nylon loops,
soaked in cryo-protectant solution, and then mounted on the goniometer head of X-ray diffraction
equipment for structural analysis. However, crystallographers often wrestle with some problem during
these operations, because protein crystals are very soft and fragile. Furthermore, the operations have to be
performed manually under microscopic observation. It is very difficult to perform these operations without
damaging the crystals. To overcome these problems, we developed a new method to grow protein crystals
in high-concentration hydrogels (>1%), which are completely gelated and exhibits high-strength [1-4].
This technique offers the practical advantages of efficient protection by the hydrogel surrounding the
protein crystals, allowing them to be handled and transported without affecting any later crystallographic
analysis. Indeed, some protein crystal grown in hydrogel reflects the highest resolution of those reported in
the Protein Data Bank. The crystal process can also be directly applied to hydrogel-grown crystals using a
femtosecond laser without mechanical damage to the crystals (Fig. 1) [5,6]. The X-ray diffraction data from
these crystals are suitable for structure analysis. Additionally, we indicated that the major advantages of this
technique are increased the number of protein crystals [7,8] and the mechanical stability of the crystals
while considerably reducing the osmotic shock [8,9].
Fig. 1: Photographs of lysozyme crystals
grown in agarose gel before and after
processing. (a, d) Crystals before processing.
(b, e) Crystals after processing. (c, f) The
excess of hydrogel surrounding the crystals
was carefully removed with Microknife and
CrystalRemover tools (SOSHO Inc.).
References
[1] Sugiyama S, et al., Jpn. J. Appl. Phys. 48, No.105502 (2009)
[2] Yoshikawa HY, et al., J. Cryst. Growth 311, 956-959 (2009)
[3] Matsumura H, et al., J. Synchrotron Rad. 18, 16-19 (2011)
[6] Hasenaka H, et al., J. Crystal Growth 312, 73-78 (2009)
[7] Tanabe K, et al., Appl. Phys. Express 2, No.125501 (2009)
[8] Sugiyama S, et al., Cryst. Growth Des. 13, 1899-1904 (2013)
[9] Sugiyama S, et al., J. Am. Chem. Soc 134, 5786-5789 (2012)
64
Abstract
Non-conventional techniques for protein crystallization and for
separating nucleation and crystal growth processes
Moreno A
Instituto de Quimica, Universidad Nacional Autonoma de Mexico. Av. Universidad 3000.
Colonia UNAM Mexico D.F. 04510, MEXICO.
New strategies are suggested for protein crystallization either in solution-growth or in gelgrowth by using different crystal growth devices applying electric and magnetic fields. The
effect of combining both electric and magnetic fields is shown and is reviewed. Proteins
with differing proportions of α-helices and β-sheets, and crystallized in different
crystallographic space groups are studied. The crystal quality is improved by using both
an electric field as well as a strong magnetic field to orient protein molecules, and gelgrowth (high concentrations of agar) to control the transport phenomena and crystal
growth. Some advantages to increase the crystal size by using temperature gradients are
also shown for a model protein (Glucose Isomerase). The crystal quality enhancement for
crystals having marginal conditions for X-ray diffraction is discussed. Finally, it is
suggested that in order to separate the nucleation and crystal growth processes by using
these electromagnetic fields or temperature variations will help to obtain either large
amount of small crystals to perform Protein Powder X-ray Diffraction or big single crystals
to perform the classic Single Crystal Protein X-ray Crystallography (or for the Protein
Neutron Diffraction Crystallography).
Acknowledgements: The author (A.M.) gratefully acknowledges financial support from
CONACYT-Mexico Project No. 175924.
65
Abstract
Improving protein crystal quality using diamagnetic levitation
containerless method
Cao HL1, Sun LH2, Li J2, Tang L2, Lu HM1, Guo YZ1, He J1, Liu YM1, Zhang CY1,
Guo WH1, Huang LJ1, Shang P1, He JH2*, Yin DC1*
1Key
Lab for Space Bioscience & Biotechnology, School of Life Sciences, Northwestern Polytechnical
University, Xi’an, Shaanxi, PR China
2Shanghai Institute of Applied Physics, Chinese Academy of Science, Shanghai, People's Republic of China
Improving protein crystal quality is an important work due to the requirement on obtaining high resolution
diffraction data for high precision structure determination in protein crystallography. However, obtaining
high quality protein crystals is often a challenge. There have been many efforts to develop methods to grow
high quality protein crystals. Among there methods, containerless crystallization techniques are considered
to be a good solution because there are no solid contacts with the grown crystals so that there is no
contamination from the solid surface, and no lattice mismatch which may cause stress and result in
deterioration of the crystal quality.
In this report, we will show an investigation of quality comparison of the crystals grown under four
conditions, including three containerless environments (diamagnetic levitation containerless condition,
agarose gel, and silicon oil) and one condition having contact with solid surface. Seven proteins were
crystallized under the four conditions, and crystal quality was assessed in terms of resolution limit, mean
mosaicity and Rmerge. Figure 1 shows a comparison of normalized resolution limit of the 7 kinds of protein
crystals grown under the four conditions.
Figure 1. A comparison of the resolution limit of protein crystals grown under different environments. The
data were averaged and normalized based on the resolution of the crystals grown in the control.
It was found that the crystals grown under the three containerless conditions demonstrated better
morphology than that of the control. X-ray diffraction data indicated that the quality of the crystals grown
under the three containerless conditions was better than that of the control. Among the three containerless
techniques, the diamagnetic levitation technique exhibited the best performance in enhancing crystal
quality. Crystals obtained from agarose gel demonstrated the second best improvementin crystal quality.
The study indicated that diamagnetic levitation technique is indeed a favorable method to grow high-quality
protein crystals, and its utilization is thus potentially useful in practical efforts to obtain well-diffracting
protein crystals.
Acknowledgements
This work was supported by the National Basic Research Program of China (973 Program) (Grant No.
2011CB710905), the National Natural Science Foundation of China (Grant Nos. 31170816, 11202167,
51201137, 31200551), the PhD programs Foundation of the Ministry of Education of China
(20116102120052, 20126102120068), and the China Postdoctoral Science Foundation (Grant No.
2013T60890), the Natural Science Basic Research Plan in Shaanxi Province of China (Grant Nos.
2012JQ3009, 2013JQ4032, 2014JM4171).
References
Cao HL et al., Acta Cryst. D 69, 1901–1910 (2013)
Lu HM et al., Rev. Sci. Instrument 79, 093903 (2008)
Yin DC et al., J. Cryst. Growth 310, 1206-1212 (2008)
66
Abstract
What X-rays cannot do: Single-particle imaging, molecular
tomography and the resolution revolution in cryo-EM
Kühlbrandt W
Max Planck Institute of Biophysics, Department of Structural Biology, Max-von-Laue Str. 3,
60438 Frankfurt am Main, Germany.
We are witnessing the break of a new era in electron cryo-microscopy (cryo-EM) for
determining the structure of protein complexes at high resolution without crystals. In
single-particle imaging, a new generation of highly sensitive direct electron detectors has
precipitated a resolution revolution. The fast readout rate of these new cameras means
that beam-induced movements, so far the most serious limiting factors in cryo-EM, can
now be overcome routinely. 3D maps of a quality that rivals that of X-ray structures at 3 Å
resolution or better can now be obtained. As an example, the 3.4 Å structure of the nickeliron hydrogen transferase Frh, a multi-enzyme complex from a methanogenic archaeon,
will be presented.
The new direct electron detectors are also having a major impact in electron cryotomography (cryo-ET). We have obtained a ~18 Å resolution map of dimeric
mitochondrial ATP synthase in situ by sub-tomogram averaging. Together with other cryoET studies of mitochondria, this provides insights into the arrangement of large
membrane protein complexes in the inner membrane cristae.
Electron crystallography of two-dimensional (2D) crystals has yielded membrane protein
structures at 3Å to 6 Å resolution for more than 2 decades. An important advantage over
x-ray crystallography is that conformational changes can be induced on the EM grid and
monitored by cryo-EM and crystallographic image processing. Two examples will be
presented: light-induced channel gating in channelrhodopsin-2, and ion-induced changes
in sodium-proton antiporters. We anticipate that the new cameras will make a significant
difference to the resolution that can be achieved with 2D crystals, which at present is
limited by crystalline order.
67
Abstract
Rules need tools / Tools need rules
Fazio VJ, Schmidt B, Peat TS, Newman J
Crystallization is often touted as being the bottleneck in structural biology, and enormous
amounts of energy and resources are expended in the quest to produce the crystals
needed. Currently, there are not many software tools available to help the laboratory
based crystallizer – tools which would allow the crystallizer to know which screens are
most like the conditions in the PDB, for example. Or tools which allow one to share screen
descriptions between database systems.
One of the reasons for this is the profound lack of standards in the field, which makes it
difficult to make tools which are generalizable. This is confounded by a lack of adoption of
standards, which is often driven by the desire to use the tools that use them. We have
started to try to break this cycle, by building tools based on local data, but then using the
rigor which our local tools have enforced upon us to develop a set of data standards which
might be more widely used.
References
Newman J et al., Australian Journal of Chemistry (2014)
Newman J et al., Crystal Growth & Design, 10(6), 2785–2792 (2010)
68
Abstract
Crystallization of membrane proteins by the lipid cubic phase or in
meso method. From solubilized protein to final structure.
Caffrey M
Membrane Structural and Functional Biology Group, Trinity College Dublin, Ireland
[email protected]
One of the primary impasses on the route that eventually leads to membrane protein
structure through to activity and function is found at the crystal production stage.
Diffraction quality crystals, with which an atomic resolution structure is sought, are
particularly difficult to prepare currently when a membrane source is used. The reason
for this lies partly in our limited ability to manipulate proteins with
hydrophobic/amphipathic surfaces that are usually enveloped with membrane lipid. More
often than not, the protein gets trapped as an intractable aggregate in its watery course
from membrane to crystal. As a result, access to the structure and thus function of tens of
thousands of membrane proteins is limited. The health consequences of this are great
given the role membrane proteins play in disease; blindness and cystic fibrosis are
examples. There exists therefore a pressing need for novel ways of producing crystals of
membrane proteins. The lipid cubic phase or in meso method is still a case in point. In this
presentation, I will briefly introduce the method and then describe how the highresolution structures of two integral membrane enzymes were determined using crystals
produced by the in meso method. The projects involved extensive screening and
optimization based on temperature, precipitant components, and mesophase hosting and
additive lipids. Heavy atom derivatization with Se-methionine and by pre-labelling, cocrystallization and soaking, as well as sulfur-SAD, were all investigated with a view to
experimental phasing.
References
Li D et al., Nature 49, 521–524.
Li D et al., Cryst. Growth Des. 14, 2034−2047.
Supported in part by Science Foundation Ireland (12/IA/1255) and the National Institutes
of Health (GM61070, GM075915).
69
Abstract
Linker-assisted crystallization of protein complexes: Application to
TIR domains
Ve T1, Williams S J, Kobe B1
1School
of Chemistry and Molecular Biosciences, University of Queensland, Brisbane,
Australia
Complexes of weakly interacting proteins are generally not very amenable to structural
studies by most structural biology approaches, including crystallography. We reasoned
that by covalently linking the interacting partners, and thereby promoting the interaction
by increasing the local concentrations of the binding partners and controlling their
stoichiometries, we can make the complexes more amenable to structural studies.
Following some successful reports in the literature, we have applied the linker strategy to
the TIR (Toll/interlukin-1 receptor) domain-containing proteins. TIR domains are found
in diverse proteins with functions in innate immunity, such as Toll-like receptors in
mammals and resistance (R) proteins in plants, as well as bacterial proteins that interfere
with host immune signaling. Self-association is key to the signaling function of TIR
domains in these pathways, but is weak by design and the stabilization of the interaction
by membrane attachment or within the context of larger complexes represents a central
regulatory mechanism within the signaling process. We show that the addition of linkers
between TIR domains facilitates the expression and purification of most of the TIR:TIR
domain complexes investigated. In some cases we found that this strategy improved the
yield of soluble protein forms, and in one case enabled the production of soluble protein
that could not be produced independently. We demonstrate that in one case, the addition
of a linker facilitated the crystallization of a TIR:TIR domain complex (Williams et al,
2014).
References
Williams SJ, et al., Science 344, 299-303 (2014)
70
Abstract
Mitochondrial inner-membrane ABC transporter, ATM1 – crucial
steps in crystallization and structure determination
Srinivasan V
LOEWE Zentrum für Synthetische Mikrobiologie SynMikro, Marburg, Germany.
Mitochondria contain several adenosine 5‘-triphosphate (ATP) binding cassette (ABC)
transporters that are crucial for organellar communication with the cytosol and nucleus.
The yeast mitochondrial ABC transporter Atm1, in concert with glutathione, functions in
the export of a substrate required for cytosolic-nuclear iron-sulfur protein biogenesis and
cellular iron regulation. Defects in the human ortholog ABCB7 cause the sideroblastic
anemia, an iron storage disease with iron-loaded mitochondria (sideroblasts), and defects
in heme metabolism. To date, structural information on eukaryotic ABC exporters is
available only for P-glycoprotein (P-gp) that is implicated in multidrug resistance (MDR)
and the mitochondrial ABCB10. This is due to the challenges involved in working with
eukaryotic membrane proteins including expression, solubilisation, purification,
crystallization, data collection and structure solution. The key steps in the elucidation of
the crystal structures of nucleotide-free and glutathione-bound form of Atm1 will be
presented (1).
Fig. 1. Crystal structure of the mitochondrial ABC transporter Atm1. (A) Illustration of the nucleotidefree Atm1 homodimer (S. cerevisiae) with monomers colored in green and red and the nucleotide-binding
domains (NBD) facing the mitochondrial matrix. The approximate positioning of the lipid bilayer is shown
by grey dashed lines. (B) The Atm1 monomer in rainbow colors depicts the 6 transmembrane helices (TM 16) and the nucleotide-binding domain (NBD).
References
1. Srinivasan V et al., Science 343, 1137-1140 (2014)
71
Abstract
Microfluidic tools for structure determination of membrane
proteins
Abdallah BG, Kupitz Ch, Fromme P, Ros A
Department of Chemistry and Biochemistry, Arizona State University, Tempe AZ, USA
X-ray free electron lasers (XFEL) have been proposed to study the structures of many important proteins
that do not readily form large crystals by enabling ‘diffraction before destruction’ with small crystals.
Important proof of principle results have been demonstrated with serial femtosecond X-ray crystallography
(SFX) using XFELs, unlocking an enormous potential for protein crystallography without the cumbersome
need to grow large crystals. However, SFX is still immature with many challenges to be overcome before it
can serve as a general approach to structure determination of proteins. For example, SFX probes protein
crystals by injecting a fine stream of crystals, which is intersected with an XFEL. The method still requires a
considerable amount of protein solution and up to several tens of thousands of diffraction patterns for data
analysis. Moreover, methods to routinely grow or fractionate protein nanocrystals are lacking.
Consequently, we are developing microfluidic methods to grow and isolate nanocrystals. We developed a
diffusion-based method establishing a gradient of crystallization conditions polling for nanocrystal growth.
This was achieved by imaging microfluidic crystal growth with second harmonic non-linear imaging of
chiral crystals (SONICC). A large parameter space in a crystallization phase diagram is probed with one
experiment, expediting the process of phase diagram development, which has been successfully applied to a
complex membrane protein, namely photosystem I. Additionally, we have created a microfluidic sorting
system driven by size-dependent dielectrophoresis to selectively sort desired nano-crystals into collection
chambers. We have demonstrated the functioning principle of this device by experimentally sorting 90 and
900 nm polystyrene beads in excellent agreement with numerical models. In addition, photosystem I
crystals were sorted into 100 nm sized fractions, as confirmed by dynamic light scattering and particle
tracking measurements. We are now moving into the next phase of these projects to improve the
throughput of nanocrystal sorting to flow rates compatible with current injector technology for SFX. We
have also designed new ‘split-and-recombine’ microfluidic devices to achieve pL-nL multiplexed batch
crystallization combined with SONICC.
Fluorescence microscopy image of the ‘heart’ of a microfluidic sorter capable of separating nano- from
microcrystals of photosystem I. Larger crystals are focused in the right center outlet channel, whereas
smaller particles are deflected into side channels. The scale bar is 50 µm.
Abdallah BG et al., ACS Nano 7, 9129-9137 (2013)
Abdallah BG et al., ACS Nano 7, 10534–10543 (2013)
72
Abstract
G protein-coupled receptors: Structure, function and drug
development
Tate CG
MRC Laboratory of Molecular Biology, Hills Road, Cambridge CB2 0QH, UK
[email protected]
Structural studies of G protein-coupled receptors (GPCRs) are hampered by their lack of
stability in detergents and their conformational flexibility. We have developed a mutagenic
strategy combined with a radioligand binding assay to isolate thermostable mutants of
GPCRs biased towards specific conformations. This has allowed us to determine the
structures of the b1-adrenergic receptor, adenosine A2A receptor and neurotensin receptor
bound to either agonists, partial agonists, inverse agonists or biased agonists. I will
discuss the strategies used for thermostabilisation and some highlights from the
structures determined. In addition, I will discuss how the thermostabilisation platform
has been used by the spin-out company, Heptares Therapeutics, to develop novel
compounds for the treatment of neurological disorders that will be entering clinical trials
this year.
73
Poster presentations
The „Alster“
74
Poster presentations
The poster presentations will take place in the ground floor of the
main conference building (Edmund-Siemers-Allee 1) close to the
main entrance. See the image below with the detailed arrangement.
The poster boards are arranged and labeled numerically. Please
check your personal poster number next to your abstract in this
book. If you like to present an additional poster to support your
talk and a poster number has not been assigned yet (poster
number P-1XX), contact the local organisation team during the
conference. There will be remaining space on the boards. Please
also contact the local staff if you like to present an additional
poster on short notice that is not registered in this abstract book.
All material to fix the poster or additional matrial will be provided.
Why not prepare A4-handouts of your poster to give away to
interested parties? Within the first three days of the conference
there will be a total of five poster sessions for discussion.
Towards the end of the conference, poster/oral presentation prices
will be awarded (100 to 300 Euro). Catering will be offered on all
floors. Signs will guide you to WCs. As you enter the building they
are found on the right hand side.
Ground floor layout of the venue
75
Poster P-1
Abstract
Improvement of protein crystals by addition of water on
setup of vapour diffusion crystallization
Poulsen JCN1, Ørum S1, Viborg A2, Fredslund F1,2, Nakai H2, Svensson B2, Maher AH2,
Lo Leggio L1
1Department
of Chemistry, University of Copenhagen, Denmark; 2 Enzyme and Protein
Chemistry, Department of Systems Biology, Technical University of Denmark, Denmark
Even when crystals of a new protein can be obtained from screening, and despite the
improvement in microfocus beamlines allowing data collection from ever smaller and
even multiple crystals, reproducibility and optimization of initial crystallization conditions
continue to be of great importance. We report here examples where crystallization
reproducibility and/or the quality of protein crystals, among others carbohydrate
modifying enzymes from probiotic organisms, could be improved by addition of water to
protein and reservoir on setting up of vapour diffusion crystallization.
JNP is partly financially supported by CoNeXT a University of Copenhagen interfaculty
collaborative project (Co(penhagen University Ne(utron and) X-(ray) T(echniques).
76
Poster P-2
Abstract
Novel insight into the mechanism of the amidases
Weber BW1, Kimani S1 and Sewell BT1,2
1Structural
Biology Research Unit, University of Cape Town, South Africa, 2Institute of
Disease and Molecular Medicine, UCT, Cape Town South Africa
Amidases belong to the nitrilase superfamily of enzymes and convert amides to the
corresponding carboxylic acids and ammonia. It is generally accepted that the reaction is
catalysed by three active site residues: cysteine, glutamate and lysine, the catalytic triad. It
is postulated that the glutamate acts as a general base catalyst which primes the cysteine
sulphur for nucleophilic attack on the substrate carbonyl carbon. This leads to the
formation of a tetrahedral intermediate which is stabilised by the lysine before its collapse
and the release of ammonia.
Recently we have shown the critical importance of a fourth, positionally conserved
residue, a glutamate, in the reaction mechanism of the amidase from Geobacillus Pallidus.
When this glutamate was mutated to either a leucine or an aspartate the enzyme was
rendered completely inactive. Our results show that this “EKE” device positions the
substrate in precise stereo-electronic alignment for the nucleophilic attack by the active
site cysteine. Quantum mechanics/Molecular mechanics (QM/MM) modelling of the active
site has indicated that the hydrogen on the sulphur of the active site cysteine may be
positioned too far away from the glutamate for it to perform its proposed role as general
base catalyst. This may indicate that in order for nucleophilic attack to occur, only precise
stereo-electronic alignment by the “EKE” device is necessary. We therefore seek to test this
hypothesis by using neutron diffraction to reveal the positions of the hydrogen atoms in
the active site.
References
Pace HC and Brenner, C. Genome Biol.2, reviews0001.1–0001.9. (2001)
Kimani SW et al., Acta Crystallogr. Sect. D-Biol. Crystallogr. 63, 1048-1058 (2007)
Weber BW et al., J Biol Chem. 288(40), 28514-28523 (2013)
77
Poster P-3
Abstract
Biophysics research support at NASA/MSFC
Karr LJ
NASA/Marshall Space Flight Center, EM10/BLDG 4464, Huntsville, AL 35812
Biophysics is a developing program out of NASA’s Space Life and Physical Sciences Division. The
program consists of four areas: Biological Macromolecules, Biomaterials, Biological Physics, and
Fluids for Biology. Marshall Space Flight Center (MSFC) is responsible for technical support for
Biological Macromolecules and Biomaterials. MSFC has played a key role in Biological Crystal
Growth in Microgravity since its inception over 30 years ago. After several interruptions to the
program due to the Space Shuttles Challenger and Columbia incidents, and more recent
programmatic shifts within NASA, the program is once more active and MSFC is supporting
several Microgravity Experiments onboard the International Space Station. From these
experiments, NASA is expecting the achievement of structures from biologically important
macromolecules, including many membrane proteins. Additionally it is hoped to conclusively
show with significant numbers of macromolecules, the benefits to growing crystals under
microgravity conditions. Moreover, several experiments are planned to determine the physical
parameters behind these successes. Marshall Space Flight Center has considerable capabilities to
support investigators, including X-Ray Diffraction equipment, Biochemical and Molecular Biology
equipment, and equipment for measuring crystal face growth rates and the solubility of
macromolecules at various temperatures.
A new area presently being evaluated for inclusion in the Biophysics Program at MSFC, is
Biomaterials. A workshop in Biomaterials was recently held as part of the broader scope of a
NASA-sponsored MaterialsLAB Workshop, April 15-16, 2014, in Arlington, Virginia. Many
responses to a Request for Information call were discussed at the workshop. Because of the
interest shown by the research community, another workshop focused solely on Biomaterials will
be held within the Materials Research Society Fall Meeting, November 30 – December 5, 2014, in
order to reach out an even broader group of researchers.
78
Poster P-4
Abstract
Elucidating structural mechanics of membrane proteins by time
resolved serial femtosecond crystallography
Coe J1, Kupitz C1, Conrad C1, Roy-Chowdhury S1, Zatsepin N2, Basu S1, Perera S3,
Chawla U3, Vaughn M1, Brown M3, Fromme P1
1Department
of Chemistry and Biochemistry, Arizona State University, Tempe, Arizona, USA; 2 Department of Physics,
Arizona State University, Tempe, Arizona, USA; 3Department of Chemistry and Biochemistry, University of Arizona,
Tucson, Arizona
Serial femtosecond X-ray crystallography (SFX) is a recently developed technique in which a stream on nanomicrocrystals are subjugated to an extremely brilliant femtosecond pulse of X-rays, generated by a free electron laser
(FEL), that allow diffraction before the ensuing Coulomb explosion destroys the crystal (Fig. 1). The use of unique
crystals for each diffraction pattern allows time-resolved studies of fast and irreversible processes such intermediate
states and substrate dissociation. This can be achieved for photoactive proteins by introducing a photoexcitation to the
protein stream at a specified time, τ, prior to interaction with the FEL, resulting in a populated transition state being
probed. The technique allows the intermediate states of important biological reactions, such as the electron transport
and subsequent undocking of ferredoxin (Fd) from photosystem I (PSI) and the trans-cis isomerization of retinol in
rhodopsin, to be studied.4
PSI is a membrane protein complex that plays an integral role in photosynthesis, catalyzing the terminal step
of photo-excited electron transport through the thylakoid membrane. This process results in the PSI bound protein Fd
becoming reduced and undocking to participate in the reduction of NADP+ to NADPH, which is crucial in the subsequent
conversion of CO2 into carbohydrates. The exact molecular mechanism by which Fd is reduced and successively
undocks from PSI is unknown, although spectroscopic studies and preliminary SFX work have provided evidence that it
is a multistep process with transition states that occur on the orders of nano and microseconds.1,2 Revealing the
electron transfer mechanism between PSI and Fd will be a significant step toward understanding of the full
photosynthetic mechanism as well as improving our grasp of electron transfer within large membrane complexes.
Rhodopsin is another membrane protein, specifically a G-protein-coupled receptor, that provides the basis for
vision in many animals. Rhodopsin is activated through photoexcitation that leads to an irreversible reaction in which
the cofactor retinal, bound through a Schiff base linkage, undergoes isomerization from 11-cis-retinal to all-transretinal. This process proceeds through multiple intermediates, resulting in metarhodopsin II in which the Shiff base is
deprotonated and is no longer covalently bound. The metarhodopsin II intermediate is the active state of rhodopsin
that gives rise to the ensuing cascade chemical reaction that triggers the visual process. While there exist
crystallographic structures for the dark and most of the intermediate states, there is currently active debate as to the
structure of metarhodopsin II since conclusions drawn from individual experiments have led to contradictory results.3,4
There is speculation that these contradictions may stem from selective radiation damage near the retinal. The aim of
this research is to uncover the mechanism of electron transfer between PSI and Fd and to establish undamaged
structures of intermediates in the photolysis of rhodopsin, particularly metarhodopsin II.
Time-resolved SFX experimental setup
Figure 1: A fully hydrated jet of protein in its
mother liquor at ambient temperature is first
photoexcited by an optical pump probe at a time
τ to populate an excited state. This is followed
by nteraction with the high flux XFEL beam,
operating in femtosecond pulses, which results
in diffraction of the undamaged crystal prior to
its complete destruction, the basis of the
"diffract before destroy“ principle. Diffraction
patterns are then recorded on the CSPAD
detector for subsequent data analysis. 1,5
References
A et al., Optics Express 20.3, 2706-2716 (2012)
2Setif P Biochim. Biophys. Act 1507.1, 161-179 (2001)
3Choe H-W et al., Nature 470, 651-655 (2011)
4Standfuss J et al., Nature 471, 656-660 (2011)
5Neutze R et al., Nature 406, 752-757 (2000)
1Aquila
79
Poster P-5
Abstract
Dissolution behaviour of lysozyme crystals
Ferreira C1, Damas AM2, 3, Martins PM1, 2, Rocha F1
1
LEPABE, Departamento de Engenharia Química, Faculdade de Engenharia da Universidade do Porto, Rua
Dr. Roberto Frias, 4200-465 Porto, Portugal; 2 ICBAS – Instituto de Ciências Biomédicas Abel Salazar,
Universidade do Porto, Largo Prof. Abel Salazar, 2, 4099-003 Porto, Portugal; 3 IBMC – Instituto de Biologia
Molecular e Celular, Rua do Campo Alegre, 823, 4150-180 Porto, Portugal.
In the present work, the dissolution of lysozyme crystals under microbatch conditions was studied. This
study has arisen from the need to correctly interpret the behavior of this system in nucleation experiments
(Galkin & Vekilov 2001). A drop of lysozyme solution was conducted to a high supersaturation by
decreasing the temperature, and after the occurrence of nucleation the system was heated to a temperature
value allowing the complete dissolution of the crystals formed. Then, the system was cooled again to
promote nucleation. This process was repeated continuously using the same system (Teychené & Biscans,
2012). It was observed that at undersaturation conditions, lysozyme crystals formed did not dissolve
completely. At early stages, the dissolution of crystals was complete; nevertheless, as the number of cycles
increased, the crystal dissolution was less extensive, until, in some cases, to block. Also, it was verified that,
as the experiments were repeated continuously, this phenomenon occurred mainly for drops of smaller
volume (see Figure 1).
Figure 1: Dissolution behaviour of lysozyme crystals at 37 ᵒC, in drops of 0.7 µL (triangles) and 0.2 µL
(squares) of solution with 25 mg/mL of lysozyme, 3% (w/v) of NaCl and 0.2 M of sodium acetate at pH 4.7
References
Galkin O & Vekilov PG, Journal of Crystal Growth 232, 63–76 (2001)
Teychené S & Biscans B, Chemical Engineering Science 77, 242-248 (2012)
80
Poster P-6
Abstract
New sample delivery methods for serial femtosecond
crystallography
Conrad C1, Kupitz C1, James D2, Wang D2, Plentnev S3, Weierstall U2, Spence J2,
Fromme P1
1Department
of Biochemistry, Arizona State University, Tempe AZ; 2Department of Physics, Arizona State
University, Tempe, AZ; 3National Caner Institute, SAIC-Fred, Argonne IL
Serial femtosecond crystallography (SFX) is a relatively new structural biology technique that is based on
femtosecond pulse X-ray diffraction data via X-ray free electron lasers (X-FEL), allowing for challenging
protein structures to be solved from tiny crystals at room temperature1. In the SFX technique microcrystals
are delivered in a liquid jet to an X-FEL, producing a diffraction pattern immediately before the crystals are
destroyed, resulting in minimal radiation damage.2 Technique development of sample delivery for SFX aims
to enhance the technique by allowing the ability for fast mixing in the liquid jet and the prospect of smaller
sample volumes. Because samples are not immobilized or frozen in SFX, determination of induced spatial
and temporal conformational changes “on the fly” is possible. Fast mixing of two liquids, such as those
containing an enzyme and a substrate, immediately before being exposed to the X-ray beam would allow
conformational changes to be seen by snapshot diffraction.3 As a model for the liquid mixing jet, mKate, a
protein that undergoes a pH dependent conformational change was chosen (Fig 1).4 Crystallization
conditions to produce mKate microcrystals have been established, however, the time delay necessary for the
conformational change must be known. To determine this τ, fluorescent correlation spectroscopy in
conjunction with fluid dynamic modeling is being employed. The second technique under development is to
mimic the highly desirable viscous properties of lipidic cubic phase (LCP).5 LCP has been a successful
delivery method for membrane proteins, though, LCP is not suitable as a general delivery system. Most SFX
experiments have relied on a liquid jet, delivering sample in the mother liquor focused into a stream by
compressed gas.6 However, this liquid stream moves at a fast rate only allowing approximately every 1 out
of 10,000 crystals to be probed by the X-ray beam, meaning that most of the valuable sample is wasted.
Sample delivered in an inert, viscous media with similar properties to LCP would allow a reduction of
sample consumption by a factor of 10-50. This would allow for SFX data collection on many new proteins,
which are currently sample limited. Methods of mixing crystals with the viscous media and growing crystals
within the viscous media are being explored. This work aims to further develop SFX by use of mixinginduced time resolved measurements for time resolved data collection and an inert, viscous media that
would allow for sample limited protein crystals that are incompatible with LCP to be probed by SFX.
Conformations of the mKate Chromophore
Figure 1: At low pH the chromophore
is predominately in the trans
conformation and no fluorescence is
seen but at high pH the chromophore
adopts a cis conformation resulting
in high fluorescence.4
Trans
Cis
Chapman H et al., Nature, 470, 7332(2011)
Neutze R et al. Nature 406, 752-757 (2000)
Aquilla A et al., Optics Express, 3, 2706-2716 (2012)
Pletnev S et al., J. Biological Chem. 283, 28980-28987 (2008)
Liu W et al., Science 342, 1521-1524 (2013)
DePonte D et al., J. Phys. D. Appl. Phys. 41, 195505 (2008)
81
Poster P-7
Abstract
Engineering the ATP synthase rotor: Customized ion-to-ATP ratios
and ion specificity
Pogoryelov D1, 2, Krasnoselska OG1, Klyszejko AL1, Leone V1, Faraldo-Gómez JD1,
Meier T1
1Max-Planck
Institute of Biophysics, Max von Laue str. 3, Frankfurt am Main, DE-60438,
Germany;
of Biochemistry, Goethe University of Frankfurt, Frankfurt am Main,
DE-60438, Germany
2Institute
E-mail: [email protected]
The F1Fo-ATP synthase membrane-embedded rotor consists of an oligomeric ring of csubunits (c-ring), of which each subunit provides one ion binding site and contributes to
the ion specificity of the enzyme. We addressed the question about how the ATP
synthase’s "ion-to-ATP ratio" could be manipulated. To do so, we investigated the
influence of a conserved stretch of glycines at the c/c-subunit contacts in the Ilyobacter
tartaricus c11 ring by site-directed mutagenesis. Structural and biochemical experiments
show a direct influence of selected mutations on the c-subunit stoichiometry, revealing c1016 rings. Protein interaction studies demonstrate that the assembly of the F1Fo rotor
complex is independent of the c-ring size and stoichiometry. Notably, the mutant ATP
synthase harboring the larger c12 instead of c11 ring synthesizes ATP at a lower membrane
potential threshold – that is, at higher ion-to-ATP ratio. We also tackled the question about
how the Na+ vs. H+ ion selectivity of the Fo rotor could be manipulated. We produced a set
of I. tartaricus c-ring mutants with substitutions of the critical residues at the ion-binding
site. To probe the different ion-binding properties of these mutants we measured the ATP
synthesis rates and the effect of the Fo-inhibitor dicyclohexylcarbodiimide (DCCD) under
variable pH and salt conditions. Our results suggest that the inherent Na+ selectivity of the
c11 ring can be adjusted by specific point mutations in the c-subunit in a way that the
selectivity of the binding site is significantly enhanced for H+ against Na+ under
physiological conditions. The work provides the exciting perspective to design and
engineer functional ATP synthases with customized ion-to-ATP ratios and ion specificity.
References
Pogoryelov D et al., Proc Natl Acad Sci USA 109, E1599-1608 (2012)
Pogoryelov D et al., Nat Chem Biol, 6(12), 891-899 (2010)
Krah A, Pogoryelov D et al., Biochim Biophys Acta 1797(6-7), 763-772 (2010)
Krah A, Pogoryelov D et al., J Mol Biol 395, 20-27 (2010)
Pogoryelov D et al., Nat Struct Mol Biol 16, 1068-1073 (2009)
Meier T et al., J Mol Biol 391, 498-507 (2009)
82
Poster P-8
Abstract
TupA and ModA: Characterization of tungstate and molybdate
binding proteins from Desulfovibrio alaskensis G20
Cordeiro RSC1, Otrelo-Cardoso AR1, Nair RR1, Correia MAS1, Santos-Silva T1 and Rivas
MG1,2
1REQUIMTE/CQFB, Departamento de Química, Faculdade de Ciências e Tecnologia, Universidade Nova de
Lisboa, Portugal
2Departamento de Física, Facultad de Bioquímica y Ciencias Biológicas, Universidad Nacional del Litoral,
Argentina
Email of corresponding author: [email protected]
Keywords: TupA, ModA, Tungsten, Molybdenum, Biochemical characterization, Crystallization, Diffraction
Molybdenum (Mo) and tungsten (W) are chemically analogous elements that are found in the environment.1
In biological systems, Mo and W are incorporated into the active site of enzymes through binding to a
pyranopterin moiety.2 The Mo and W enzymes have an important physiological role, including the catalysis
of some main reactions in the metabolism of carbon, nitrogen and sulfur.3 These elements are incorporated
into the cell through specific transport systems, one of them known as TupABC and ModABC. This is
composed of a protein located in periplasm (TupA/ModA), a membrane protein (TupB/ModB) and a protein
confined in the cytoplasm (TupC/ModC) responsible for the hydrolysis of ATP, thus generating the energy
needed to transport the oxyanions to the cell.4
TupA from Desulfovibrio alaskensis G20 has been successfully crystallized using PEG3350 as precipitating
agent and a complete dataset was collected at ID 23-1 beamline of the ESRF (European Synchrotron
Radiation Facility). The crystals diffracted up to 1.4 Å resolution and belong to the P21 space group with the
cell constants a=52.25 Å, b=42.50 Å, c=54.71Å and =95.43˚.5 On the other hand, ModA, from Desulfovibrio
alaskensis G20 was also crystallized using PEG6K as precipitating agent and a complete dataset was
collected at ID I03 beamline of Diamond (Diamond Light Source). The crystals diffracted up to 2 Å resolution
and belong to space group C2, and the dataset is currently being processed. In order to understand the
binding mode and the metal specificity of the protein TupA, site directed mutagenesis has been carried out
and three different mutants have been designed, substituting arginine 118 to a lysine (R118K), a glutamine
(R118Q) and a glutamate (R118E). Native polyacrylamide gel electrophoresis was performed for a
qualitative analysis of binding with tungstate and molybdate for native TupA and the three mutations as
well as for the native ModA. In addition, urea-polyacrylamide gel electrophoresis was used to see the
conformational behavior of each protein, in presence and absence of ligands as well. In order to
quantitatively measure the binding between the proteins and ligands and determine de dissociation
constant (KD), isothermal titration calorimetry (ITC) was also performed. Crystals of the mutated forms of
TupA are being prepared in the presence and absence of ligands as well as crystals of native TupA and ModA
in presence of both tungstate and molybdate.
The structural and biochemical characterization of the proteins involved in ion transport will contribute to
the understanding of the specificity of the metal uptake process and its effect in the cell.
References
1Smart
JP et al., Mol Microbiol, 74, 742-757 (2009)
G et al., Molecular Microbiology of Heavy Metals, 6, 423-451 (2007)
3Aguilar-Barajas E et al, Biometals, 24, 687-77 (2011)
4Mota S et al., Bacteriol, 193, 2917-2923 (2011)
5Otrelo-Cardoso AR et al., Int. J. Mol. Sci, 15, 1-17 (2014)
2Schwarz
83
Poster P-9
Abstract
Structural characterization of PROPPINs
Scacioc A1,2, Busse RA1,2, Kühnel K1
1Max
Planck Institute for Biophysical Chemistry, Göttingen, Germany; 2The Göttingen
Graduate School for Neurosciences, Biophysics, and Molecular Biosciences, Georg-August
University Göttingen, Göttingen, Germany
PROPPINs
-propellers that bind polyphosphoinositides) are a family of
phosphoinositides binding
proteins that are important in autophagy. They are peripheral membrane proteins that
have two binding pockets that specifically bind PI3P and PI(3,5)P2. There are three
PROPPIN paralogs involved in yeast autophagy: Atg18, Atg21 and Hsv2 (homologous with
swollen vacuole phenotype 2). The structure of Kluyveromyces lactis Hsv2 was determined
in our group (Krick et al). Trials for structure
determination were done on other PROPPINs from different organisms. However, due to
the long loops
present, especially loop CD in blade 6, no success was recorded. Loop 6CD is important for
membrane
binding. Needles diffracting at 7.7 Å were observed for Pichia angusta Atg18. Using in situ
proteolysis, we have obtained crystals shaped as plates. From these we determined the
structure this homolog at 1.8 Å resolution. Loop 6CD, consisting by length, one third of the
protein, was not present in the structure. Interestingly, several hits were obtained for
different conditions in the presence of phosphates. In the final structure, a phosphate is
bound to binding site 2. However, binding site 1 has one of the molecules from the
crystallization condition. In order to identify the molecule out of several potential
candidates, before freezing, we soaked the crystals with the crystallization condition
minus one of the components in presence of excess phosphate. Finally, a structure with
both binding sites occupied by phosphates was determined at 2.5 Å and the molecule
present in the 1.8 Å structure was identified.
References
Krik R et al., Proc Natl Acad Sci U S A. 109(30), E2042-9 (2012)
84
Poster P-10
Abstract
Structural and functional studies on nitric oxide synthase
complexed with Re(CO)3- compounds
Leisico F1, Correia MAS1, Morais M2, Oliveira BL2, Macedo AL1, Romão MJ1, SantosSilva T1, Correia JDG2
1UCIBIO@REQUIMTE,
Dep. de Química, Faculdade de Ciências e Tecnologia, Universidade Nova de Lisboa,
FCT-UNL, 2829-516 Caparica, Portugal; 2Unidade de Ciências Químicas e Radiofarmacêuticas, IST/ITN,
Instituto Superior Técnico, Universidade Técnica de Lisboa, Estrada Nacional 10, 2686-953 Sacavém,
Portugal
Nitric oxide synthase (NOS) catalyzes the O2-dependent conversion of L-arginine to L-citruline and nitric oxide (NO).
The enzyme presents three isoforms, namely neuronal (nNOS), endothelial (eNOS) and inducible (iNOS), in which nNOS
and eNOS are constitutively expressed and Ca2+-dependent while iNOS is inducible expressed and Ca2+-independent. NO
is a key physiological modulator in mammals which, among other functions, stimulates vasodilatation in the
cardiovascular system and serves as a neurotransmitter in the central nervous system. Besides mediating several
physiological functions, NO overproduction by nNOS and iNOS has been associated to several clinical disorders
including stroke, Alzheimer’s and Parkinson’s disease and cancer [1].
Monitoring NOS expression in vivo represents a promising approach to early detection of diverse
pathologies which represents a potential advantage for the treatment of such disorders. Aiming the design of innovative
radioactive probes for in vivo targeting of NOS, a set of 99mTc(CO)3-complexes containing NOS-recognizing units (e.g. LArginine derivatives, guanidine or S-methylisothiourea moieties) were established and Re(CO)3-complex analogs were
studied as “non-radioactive” surrogates [2]. Interestingly, previous results have showed that some of the rhenium
complexes have high inhibitory activity towards iNOS, namely Re1 and Re2 (Figure 1) which were characterized by KI’s
of 84 µM and 6 µM, respectively. Additionally, both Re complexes permeate through macrophage cell membranes and
interact with the cytosolic target enzyme, inhibiting NO biosynthesis in LPS-induced macrophages [3].
Aiming to structurally characterize the interaction of the synthesized metal compounds with the
proteins of interest, the oxygenase domain of human NOS of each isoform was produced. The expression system of iNOS
has been optimized and pure active enzyme has been obtained. More than a thousand crystallization conditions have
already been tested to try to crystallize the protein-metal complexes and some positive hits were obtained and multiple
crystals are under optimization using soaking and co-crystallization approaches. Several crystallization trials involving
eNOS and nNOS are being performed in order to find suitable conditions to crystallize all isoforms of NOS and to carry
out structural studies regarding the complexes of NOS proteins and the organometallic compounds.
Re1 (n=1) and Re2 (n=4)
Figure 1- Organometallic Re(CO)3
complexes which showed higest
inibitory action towards NOS
isoforms [2].
Complementary studies have also been conducted. Isothermal Titration
Calorimetry (ITC) was used to test the activity of the produced isoforms
showing that iNOS and eNOS are active in the presence of tetrahydrobiopterin
(BH4) and L-arginine. The interaction of the protein with the two Re compounds
is going to be characterized by this technique in the near future. Furthermore,
interaction of iNOS and Re1 and Re2 compounds was studied through NMR
saturation-transfer difference (STD). We identified the nearest interacting
protons in binding moieties for each compound towards to the protein,
confirming previous molecular docking studies [4]. The results will allow to shed
light on selectivity and affinity properties of these compounds toward NOS,
which constitute important interaction evidences for the design of new specific
compounds to target NOS proteins. Further crystallography and ITC analysis
will complement the all spectra of structural and functional results regarding
the three isoforms of NOS and the most interesting organometallic complexes.
References
1 Alderton WK & Knowles, Biochemistry Journal 357, 593-615 (2001)
2 Oliveira BL, et al., Journal of Organometallic Chemistry 696, 1057-1065 (2011)
3 Oliveira BL, et al., Bioconjugate Chemistry 21, 2168-2172 (2010)
4 Oliveira BL et al., Journal of Molecular Graphics and Modelling 45, 13-25
(2013)
85
Poster P-11
Abstract
Characterization of ATP-dependent transport via biophysical
analysis of the bacterial ABC exporter MsbA
Müller A1, Syberg F1, Suveyzdis Y1, Kock K 2, Schartner J1, Herrmann C2, Kötting C1,
Gerwert K1, Hofmann E1
1Department
of Biophysics, Ruhr-University Bochum; 2Department of Physical Chemistry
I, Ruhr-University Bochum
ABC transporters form a ubiquitous family of integral transmembrane proteins
responsible for the active transport of a wide variety of compounds across biological
membranes. They share a common overall architecture consisting of two transmembrane
domains (TMD), found to be responsible for substrate (allocrite) specificity, and two
nucleotide binding domains (NBD), which power transport by ATP hydrolysis. We are
working on the homodimeric E. coli ABC-Transporter MsbA located in the cytoplasmatic
membrane of Gram-negative bacteria. It is essential for translocation of Lipid A from the
inner to the outer membrane leaflet (1, 2). MsbA exports similar xenobiotics as human PGlycoprotein or other bacterial multidrug transporter like LmrA from Lactococcus lactis
(3) and therefore can be used as a prokaryotic model for eukaryotic ABC-multidrug
exporters. Despite the fact, that the X-ray structure of MsbA and other ABC-transporters
were published over the last years, the molecular details of the mechanism are poorly
understood (4).
In a first step we applied time-resolved Fourier Transform Infrared Spectroscopy (trFTIR)
to investigate isolated NBDs. We employed a cysteine-free MsbA-NBD mutant to obtain the
first time-resolved FTIR-spectra of the ATP hydrolysis reaction of an ABC transporter. This
allowed us to kinetically follow the catalytic cycle of the NBD and determine two rates
which characterize ATP binding and hydrolysis. We could identify a substrate-enzyme
complex intermediate. We also show that hydrolysis is the rate limiting step in this cycle
(5). By comparing trFTIR and Xray structures in different states of the transport cycle,
using different mutants, we want to elucidate the involved amino acids.
Benefiting from the experience with the NBD, we want to apply the same methods on the
full length MsbA protein, obtaining detailed information of the mechanisms of coupling
the ATP-hydrolysis to movement of the TMD of ABC transporters. ATR-FTIR spectroscopy
allows the investigation of hardly soluble transmembrane proteins. First results show the
successful immobilization of the full length transporter onto a germanium ATR-FTIR
crystal. This approach is expected to be applicable to other ABC-transporters. In order to
gain high resolution structures for deeper insight into the whole mechanism, we also work
on the crystallization of full length MsbA.
(1) Doerrler WT et al., J. Biol. Chem. 276, 11461 (2001)
(2) Eckford PDW and Sharom FJ, Biochem. J. 429, 195 (2010)
(3) Reuter G et al., J. Biol. Chem. 278, 35193 (2003)
(4) Ward A et al., Proc. Natl. Acad. Sci. U.S.A. 104, 19005 (2007)
(5) Syberg F et al., J. Biol. Chem. 287, 23923 (2012)
86
Poster P-12
Abstract
Novel crystal handlings of photosystem II: Characterization using
near-infrared imaging, dehydration of the glue coating crystals
using humidity gas and Laser-processing for cylinder-shaped
crystals
Umena Y1,2, Baba S3, Kawano Y4, Kumasaka T3, Kamiya N1,5
1
The OCU Advanced Research Institute for Natural Science and Technology (OCARINA), Osaka City
University, Osaka, Japan, 2 Precursory Research for Embryonic Science and Technology (PRESTO), Japan
Science and Technology Agency (JST), Tokyo, Japan, 3 JASRI/SPring-8, Research ＆Utilization Division,
Hyogo, Japan 4 RIKEN/SPring-8 Center, Advanced Photon Technology Division, Hyogo, Japan, 5 Graduate
School of Science, Osaka City University, Osaka, Japan
Molecular oxygen on Earth is generated from photosynthesis by cyanobacteria, algae and plants. Photosystem II (PSII)
catalyzes light-induced water splitting reaction leading to the production of protons, electrons and molecular oxygen in
photosynthesis. Cyanobacterial PSII is a multi-subunits membrane protein complex composed of 17 membranespanning subunits, 3 membrane-extrinsic subunits and about 80 co-factor molecules with a total molecular weight of
350 kDa as a monomer. The catalytic center of oxygen evolving complex (OEC) in PSII is composed of four Mn atoms and
one Ca atom organized in a Mn4CaO5-cluster. We reported the PSII structure at 1.9 Å resolution prepared from
Thermosynechococcus vulcanus (PDB code: 3ARC)[1].
But there are still some difficulties to prepare the good-quality PSII crystals at high reproducibility. In the observation
of PSII crystals, it is difficult to detect the problems on the crystal surface: cracks, chips and attached small crystals. This
is because dark-colored PSII includes many pigments, chlorophylls and beta-carotenes, which absorb visible light. In the
X-ray diffraction procedures, the post-crystallization treatment by gradual dehydration is essential to collect the
diffraction data at atomic resolution. To control the speed of crystal shrinking is important to keep the diffraction
quality. The isomorphism among the crystals is important in the data collection using multi-crystal averaging method.
The internal isomorphism is particularly sensitive to the data evaluation when these data are collected using soft X-ray
source.
In our study, we attempted to solve these problems of PSII crystal analysis using following novel approaches. First one
is that we developed the near-infrared imaging system for dark-colored PSII crystal. This imaging system is easy to
detect the internal damage as well as surface cracks (Fig.1). Secondly we utilized the controlled humidity gas system for
the crystal shrinking, which were established in the synchrotron facility SPring-8 in Japan and used the glue-coating
method for gradual dehydration [2]. We found improvement conditions of shrinking PSII crystal. Finally, we processed
from an angular PSII crystal to a cylinder-shaped crystal using deep-UV pulsed laser [3]. These uniform shaped crystals
were suitable for collecting the diffraction data using soft X-ray or by mult-crystal averaging method. We will show
these novel post-crystallization approaches of PSII crystal analysis.
Fig.1 PSII crystal images using visible light (A) and
using near-infrared light system (B)
Fig.2 Dehydration process of a glue-coated
PSII crystal (left) and the improved
diffraction patterns from the crystal by the
humidity changing from 96 %RH to 82 %RH
(right)
References
[1] Umena Y, Kawakami K, Shen JR, Kamiya N, Nature 473, 55-60, (2011)
[2] Baba S et al., Acta Cryst. D60, 1839–1849, (2013)
[3] Kitano H. et al., Jpn. J. Appl. Phys. 44 , L54, (2004)
87
Poster P-13
Abstract
Cloning, expression and purification of binary protein complexes
involved in lipolysis
Viertlmayr R1, Cerk IK1, Eder M1, Böszörmenyi A1, Nagy HM1, Das KMP1, Bird L2, Rada
H2, Owens R2, Oberer M1
1 Institute
2 Oxford
of Molecular Biosciences (IMB), University of Graz, Austria
Protein Production Facility UK (OPPF), University of Oxford, United Kingdom
In recent years, lipolysis has gained enormous scientific and societal interest due to the increasing
number of overweight and obesity in the population. Reasons for overweight and obesity are
mainly an imbalance between energy intake and expenditure. The so-called obesity epidemic is
associated with multiple diseases including type 2 diabetes, cardiovascular diseases, and some
forms of cancer (Greenberg AS, et al., 2006)., Excess of dietary fatty acids are stored in form of
triglycerides in adipose tissue. ATGL is the rate-limiting enzyme in lipolysis. It catalyzes the
hydrolysis of triglycerides stored in lipid droplets to diglycerides and a molecule of free fatty acid
(Lass A, et al., 2011).
The objective of this ambitious project is to characterize the structure of mammalian adipose
triglyceride lipase (ATGL) in complex with its protein inhibitor G0/G1 switch gene 2 (G0S2),
and/or its activator comparative gene identification-58 (CGI-58). Previous studies indicate that
ATGL consists of at least two domains, and it is unknown, whether the C-terminal half of ATGL
adopts typical secondary structure elements in the absence of a binding partner (Cornaciu I, et al.,
2011). Based on its small size of 11 kDa, G0S2 has a high content of hydrophobic residues and
might be unstructured by itself. Using polycistronic coexpression we tried to achieving more
stable and easy to handle protein-protein complexes.
During our work we consequently optimized construct size, tested for different expression hosts
and identified parameters which allowed us the purification of our target proteins. The proteins
and protein complexes were produced from codon-optimized genes in a polycistronic coexpression system (pST44) (Tan S, et al., 2005). We performed a high throughput approach at the
Oxford Protein Production Facility UK (OPPF) to identify the perfect vector system, fusion protein
and expression organism for our targets. Until now we have been able to overexpress all our
target proteins within the polycistronic expression system in their active form. In case of the
binary ATGL/CGI-58 and CGI-58/G0S2 complexes we screened various additives and detergents
and established a purification protocol using detergents, which increased the solubility and
stability of our fusion protein constructs.
The binary ATGL/G0S2 construct only shows suitable protein overexpression for G0S2.
Expression of ATGL seems to be inhibited by the presence of G0S2. Currently, the underlying
mechanism which inhibits ATGL overexpression is unknown. Recent progress in these ongoing
study will be shown.
References:
Greenberg AS et al., The American Journal of Clinical Nutrition 83, 461S-465S (2006)
Lass A et al., Progress in Lipid Research 50, 14-27 (2011)
Cornaciu I et al., PLoS ONE 6 (2011)
Tan S et al., Protein Expression and Purification 40, 385-395 (2005)
88
Poster P-14
Abstract
Chitinolytic enzymes of Clostridium paraputrificum J4; separation
and characterization
Dušková J1, Šimůnek J2, Tiščenko G3, Kolenko P3, Fejfarová K3, Skálová T1,
Dohnálek J1,3
1Institute
of Biotechnology, Academy of Sciences of the Czech Republic, Praha; 2Institute of
Animal Physiology and Genetics, Academy of Sciences of the Czech Republic, Praha;
3Institute of Macromolecular Chemistry, Academy of Sciences of the Czech Republic, Praha
Chitinases are hydrolases which catalyse cleavage of glycosidic bond in chitin. Chitin is
digested by certain bacteria, therefore chitinases play an essential role in their nutrition.
Higher organisms use chitinases in defence against pathogenic fungi with chitincontaining cell walls. Humans retrieve chitinases with help of Clostridium paraputrificum
J4, a human intestinal bacterium producing several extracellular chitinolytic enzymes.
This ability might even play a role in human immune defence (Zhou et al. 2004).
We have sequenced the genome of Clostridium paraputrificum J4 and discovered twelve
genes coding for diverse chitinolytic enzymes. Among them, we focused on the major 62.3
kDa chitinase Chit62J4. The enzyme possessing a four-domain structure belongs to the
glycoside hydrolase family 18. The catalytic domain has a high homology to chitinase D
from Bacillus circulans (Hsieh et al., 2010). We isolated Chit62J4 both from crude culture
medium (Dušková et al., 2011) and produced it by recombinant protein expression in
Escherichia coli. We compared the enzymatic properties of these two samples towards
diverse chitin-like substrates. We observed the highest activity towards 4-nitrophenyl
N,N’-diacetyl-β-D-chitobioside, so biosidase activity is the major activity of the enzyme.
This fact was taken into account in further studies such as pH and temperature optimum
determination, kinetic parameters determination and inhibition studies. Crystallization of
both forms is under way. So far it resulted in spherulites recognizable in UV light.
References
Dušková J et al., Acta Biochim. Pol. 58, 261-263 (2011)
Hsieh YC et al., J. Biol. Chem. 285, 31603-31615 (2010)
Zhou Zhu et al., Science 304, 1678-1682 (2004)
89
Poster P-15
Abstract
Structure of the laccase from fungi Trichaptum abietinum
Osipov EM1, Dagys M2, Casaite V2, Shleev SV13, Tikhonova TV1, Meskys R2 and
Popov VO14
1AN
Bach Institute of Biochemistry, Moscow, Russia; 2Institute of Biochemistry, Vilnius
University, Vilnius, Lithuania; 3Malmö University, Malmö, Sweden; 4RSC ‘Kurchatov
Institute’, Moscow, Russia
Laccases are members of a large family of multicopper oxidases, which oxidize a wide
range of organic and inorganic substrates accompanied by dioxygen reduction to water. An
electrode with immobilized laccase could serve either as a biocathode in biofuel cells or
biosensors for various phenolic compounds. The catalytic activity of laccase from
Trichaptum abietinum (TaL) immobilized on the electrode modified with gold
nanoparticles is threefold higher than that of laccase from Trametes hirsuta (ThL)
immobilized in similar manner.
In this work, we discuss the crystallization and structure of TaL. A comparison of the TaL
and ThL structures revealed structural differences responsible for the differences in their
catalytic activity.
Crystallization conditions were screened by the sitting-drop vapor-diffusion method.
Small needle-like clusters appeared within 2 weeks in 2 M ammonium sulfate, 0.1 M MES,
pH 6.5, and 5% 1,4-dioxane. The optimization was performed using 1.0 2.0 M ammonium
sulfate, 0.1 M MES, pH 6.5, and 1 10% 1,4-dioxane. During optimization no crystals
suitable for X-ray diffraction were obtained. To improve the quality of crystals, we used
streak-seeding. Well-shaped crystals appeared under the same conditions within 1 day
and grew to the maximum size 150x50x50 µm in one week.
The structure of TaL was determined by X-ray crystallography at 2.5 Å resolution. The
structure analysis showed that the fold of TaL is typical of all laccases and is very similar to
ThL (r.m.s.d. by C = 0.4 Å), which reveals a 59% sequence identity. The most notable
differences between two structures were found in the vicinity of the T1 site. The loops
Pro390-Asn394 and Phe159-Ala165 in the TaL structure were substituted by Pro385Ala393 and Pro160-Pro163 in ThL. We suppose that the differences in these loops affect
the arrangement of the laccase molecules on the electrode surface thus affecting the
catalytic activity of the enzymes.
This work was supported by the RFBR grant No. 13-04-40207-H.
90
Poster P-16
Abstract
Structural insights into mistranslation
Rozov A1, Demeshkina N1, Westhof E2, Yusupov M1, Yusupova G1.
1Institut
de Génétique et de Biologie Moléculaire et Cellulaire, Illkirch, France; 2Institut de
Biologie Moléculaire et Cellulaire, Université de Strasbourg, Strasbourg, France
mRNA translation by the ribosome is a complex and thus a highly error-prone process. The ribosome
demonstrates quite efficient discrimination to select a right tRNA out of the total pool, but it never reaches
similar levels of fidelity as, for example, DNA replication, with mistranslation frequency about 10-3–10-4. The
formation energy of a 3 base pair-long duplex between mRNA codon and tRNA anticodon is likely to be
sufficient to separate cognate and non-cognate tRNAs. Whereas various near-cognate tRNAs (with one
mismatch) present a challenge, since the energy level differences, low enough as they are, are additionally
influenced by the nature of the mismatch and its position in the duplex. Therefore it is safe to presume that
ribosome must utilize some kind of amplifying mechanism to bring the differences between tRNAs to an
easily discernable level.
First attempts to study the structure of ribosome decoding center were undertaken over a decade ago. The
investigation was focused on the crystal structure of the 30S subunit soaked with short analogs of mRNA
and tRNA (Ogle et al., 2002). It was suggested that ribosome undergoes certain conformational changes
induced strictly by the binding of cognate tRNA. However, this model imposed some obvious limitations on
the result. It took several more years for the crystal structures of full 70S ribosomes to reach levels attained
by the separate subunits earlier. Recently, our group has reported crystal structures of the ribosome
carrying cognate and near-cognate tRNAs in the A site (Demeshkina et al., 2012). The conformational
rearrangements of the 30S subunit appeared to be identical for all cases and, hence, non-discriminatory. The
behavior of the introduced mismatches was, however, surprising. G•U base pairs, replacing the canonical
G•C at either first or second position, adopted Watson-Crick geometry instead of expected wobble, implying
that the tautomeric equilibrium for the nitrogen bases was shifted. This finding led us to propose a new
model of the decoding process. It appears that tRNA discrimination is mediated by the rigid mold of the
ribosome decoding center. It is likely that the energy cost of fitting a mismatched base pair into the limits
imposed by the standard duplex geometry will be high enough to play a role in the rejection of near-cognate
tRNAs in favor of cognate ones.
Here we present several more structures of 70S ribosomes co-crystallized with mRNA and tRNAs featuring
various mismatches in the first and second position of the A site. Additionally, we have solved structures
with near-cognate tRNA in the P site. This structures model the rare event in which the near-cognate tRNAs
are accommodated by the ribosome and undergo translocation. The conclusions derived from our structural
investigations are corroborated by the in vivo investigation of mistranslation events (Manickam et al., 2014).
Taken together, our data substantiate the proposed model of the rigid decoding center and provide an
insight into explanation of variable protein mutation frequency.
References
Ogle JM et al., Cell, 111(5), 721-732 (2002)
Demeshkina N et al., Nature, 484(7393), 256-259 (2012)
Manickam N et al., RNA, 20(1), 9-15 (2014)
91
Poster P-17
Abstract
Conformational flexibility of the cap in the monoglyceride lipase
from Bacillus sp. H257
Aschauer P1, Rengachari S1, Schittmayer-Schantl M2, Mayer N3, Gruber K1,
Breinbauer R3, Birner-Grünberger R2, Dreveny I4, Oberer M1
1 Institute of Molecular Biosciences, University of Graz, A-8010 Graz, Austria
2 Institute of Pathology and Centre of Medical Research, A-8010 Graz, Austria
3 Institute of Organic Chemistry, Graz University of Technology, A-8010 Graz, Austria
4 School of Pharmacy, University of Nottingham, UK
Monoglyceride Lipases (MGLs) catalyze the hydrolysis of monoglycerides (MGs) into a free fatty acid and
glycerol and were described firstly in the late 1960ies. MGLs are conserved across all species yet different
biological roles in the organisms and tissues they occur are attributed to them. In mammalian lipid
metabolism, MGLs catalyze the last step of intracellular lipolysis. Additionally, they participate in signaling
mechanisms in mammalian brains hydrolyzing the endocannabinoid 2-arachidonoyl-glycerol (2-AG). Since
monoglycerides are toxic to microorganisms, MGLs also play a role in the detoxification. MGLs show high
specificity for monoglycerides (MGs). They do not hydrolyze tri- or diglycerides and do not distinguish
between different enantiomers (1-, 2- and 3-MGs). Most MGLs have varying turnover rates for MG
substrates with different chain lengths. Despite their long established importance, structural knowledge of
MGLs is only available since 2010 when the crystal structure of human MGL was determined (1-3). In 2012,
we reported the first structure of a bacterial MGL (4). The structures revealed the predicted / hydrolase
fold and an unexpected conserved architecture of the cap region compared to human MGL. This cap adopts
open and closed conformations in the human structures. In this study, we establish the structural basis for
substrate binding and specificity and by extensively varying the crystallization conditions we could collect
snapshots of the conformational plasticity of the cap region (5). We identified the key residue in these
conformational changes, established its conservation also in human MGL and established its significance for
enzyme activity.
The observed cap flexibility also provides a possible explanation for the specificity of MGLs towards
monoglycerides due to spatial restraints in substrate binding. The crystal structures of bMGL in complex
with covalently binding substrate analogues of varying chain lengths gives first insights into a steric bending
required for long chain substrates, thus suggesting a plausible explanation for decreases substrate turnover
rates. Additionally, we were successful in generating the first complex structures of an MGL with its natural
substrate 1-monolauroyl-glycerol and substrate analogs, which provides unprecedented insight into
residues involved in stabilizing the glycerol moiety at the catalytic center.
References
1. Bertrand T et al., J Mol Biol, 396(3), 663-73 (2009)
2. Labar G et al., Chembiochem, 11(2), 218-27 (2010)
3. Schalk-Hihi, C et al., Protein Sci, 20(4), 670-683 (2011)
4. Rengachari S et al., Biochim Biophys Acta, 1821(7), 1012-1021 (2012)
5. Rengachari S and Aschauer P et al., J Biol Chem, 288(43), 31093-31104 (2013)
92
Poster P-18
Abstract
The cyanotoxin microcystin: Exploring its interaction with the
regulatory protein CP12
Hackenberg C1, Dittmann E2, Lamzin V1
1 European
2 Institut
Molecular Biology Laboratory c/o DESY, Hamburg, Germany;
für Biochemie und Biologie, Universität Potsdam, Potsdam, Germany
Microcystins are the most common and notorious toxins produced by harmful cyanobacterial blooms (CyanoHABs) in
marine, brackish and fresh water habitats. They strongly inhibit protein phosphatases type 1 (PP1) and 2A (PP2A) in the
liver by covalently binding to cysteines in the active side [1,2]. This is causing severe and often lethal hepatic necrosis
and may even lead to cancer [3-5]. Since microcystins increase the resistance of CyanoHABs towards several
environmental stresses [e.g. 6], it is expected that the current climate change will further promote the proliferation of
microcystin-producing cyanobacteria [7]. Thus, microcystin-producing CyanoHABs represent a serious threat for
drinking and recreational water resources as well as human health worldwide.
Despite this importance, detailed molecular knowledge about the role of microcystin in cyanobacteria is still fairly
limited. Although it could be shown that microcystin binds in a redox-dependent manner to specific proteins in
Microcystis aeruginosa [6], still open questions remain about its binding sites, and its effect on the stability,
conformation and activity of the target proteins.
We aim to follow up on the hypothesis that it binds to the cysteine pairs of proteins and acts as protein-modulating
metabolite [6] by investigating its effect on two known target proteins: the small regulatory protein CP12 and the
photosynthetic protein phosphoribulokinase (PRK) of M. aeruginosa.
These proteins are of particular interest due to the following reasons: 1) the activity of PRK is regulated by CP12 in a
redox-dependent manner [8]; 2) CP12 is an intrinsically disordered protein with two conserved cysteine pairs forming
loops under oxidised conditions; and 3) cyanobacteria possess a variety of several CP12-variants, a.o. curiously being
fused to a CBS domain (Bateman domain) [9]. The CBS domain can be found in various organisms, where it is fused to a
wide range of proteins regulating their activity. Mutations of the CBS domain are implicated in several hereditary
human diseases [e.g. 10]. However, its function in cyanobacteria, particularly as fusion protein with the regulatory
protein CP12, is so far unknown.
The fusion of CP12 with CBS suggests a functional connection as well as different, multiple roles of CP12 in
cyanobacteria. The interaction of CP12 and PRK with microcystin may even add a further layer of complexity in the
regulation, stability and activity of these proteins in harmful cyanobacteria. Using interaction studies and
macromolecular crystallisation we aim to explore: 1) the role of the CBS domain in CP12; 2) whether CP12-CBS is still
regulating PRK; 3) the binding site of microcystin on PRK and CP12; and 4) how microcystin is affecting the stability,
conformation and activity of CP1-CBS2 and PRK.
With the results obtained we expect to significantly enhance the molecular understanding of the intracellular role and
function of microcystin as well as the ecological traits of toxic M. aeruginosa.
References
[1] Goldberg J et al., Nature 376, 745-753 (1995)
[2] Xing Y et al., Cell 127, 341-353 (2006)
[3] Lopez et al., In: Scientific Assessment of Freshwater Harmful Algae Blooms. Interagency Working Group on Harmful
Algal Blooms, Hypoxia, and Human Health of the Joint Subcommittee on Ocean Science and Technology. Washington, DC.
[4] Nishiwaki-Matsushima et al., J. Cancer Res. Clin. Oncol., 118, 420-424 (1992)
[5] Ueno et al., Carcinogenesis, 17, 1317-1321 (1996)
[6] Zilliges et al., PLoS ONE, 18, e17615 (2011)
[7] Paerl HW & Huisman J, Science, 320, 57-58 (2008)
[8] Tamoi et al., Proc. Natl. Acad. Sci. USA, 42, 504-513 (2005)
[9] Stanley et al., Plant J., 161, 824-835 (2013)
[10] Scott et al., J. Clin. Invest. 113, 274-284 (2004)
93
Poster P-19
Abstract
Structural organization and mechanism of thermostability of
histone-like HU-proteins from mycoplasma Spiroplasma meliferum
Boyko KM1,2 , Gorbacheva MA1,2, Rakitina TV1, Korgenevsky DA1, Vanyushkina AA3,
Kamashev DE3, Lipkin AV2, Popov VO1,2
1 A.N. Bach Institute of Biochemistry RAS, Moscow, Russian Federation;
2National Research Centre “Kurchatov Institute, Moscow, Russian Federation;
3SRI of Physical-Chemical Medicine, Moscow, Russian Federation
HU proteins are the most abundant DNA-binding proteins in prokaryotic organisms,
where they play substantial role in processes of DNA repair, recombination and replication
[Drlica & Rouviere-Yaniv, 1987, Kamashev et al., 2008]. Moreover, HU protein deletion
from the bacteria genome is lethal in many cases. These small (about 90 amino acids per
monomer) basic proteins occur as though homo- or hetero dimers fastened by hydrogen
bonds, salt bridges as well as large central hydrophobic core. Thus HU proteins are
convenient model for investigation of the structural basis of thermostability
[Christodoulou et al., 2003, Ramstein et al., 2003, Kawamura et al., 1998].
Mycoplasmas are the smallest known microorganisms and intercellular parasites
[Kamashev et al., 2011], which cause various mammal diseases. Up to date there was no
structural information on HU proteins from mycoplasmas. Here we present the first
structure of HU-protein from Spiroplasma meliferum (HUSpm) at 1.4A resolution.
Comprehensive structure analysis combined with DSC experiments allowed to reveal
some unique features of HUSpm and associate them with their thermal characteristics.
This work is supported by Scholarship of the President of Russian Federation for young
scientists (CP-1390.2012.4).
References
Christodoulou et al., Extremophiles 7, 111-122 (2003)
Drlica K. & Rouviere-Yaniv J, Microbiological reviews 51, 301-319 (1987)
Kamashev et al., Nucleic acids research 36, 1026-1036 (2008)
Kamashev et al., Biochemistry 50, 8692-8702 (2011)
Kawamura et al., The Journal of biological chemistry 273, 19982-19987 (1998)
Ramstein et al., J Mol Biol 331, 101-121 (2003)
94
Poster P-20
Abstract
Crystal structure of O-acetyl-L-homoserine sulfohydrolase from
bacteria Brucella melitensis causing undulant fever
Nikolaeva AY1, Boyko KM1,2, Korgenevsky DA2, Kotlov MI3, Kulikova VV3,
Revtovich SV3, Demidkina TV3, Popov VO1,2
1A.N. Bach
Institute of Biochemistry RAS
Centre “Kurchatov Institute”
3Engelhardt Institute of Molecular Biology RAS
2National Research
Enzymes that use the cofactor pyridoxal phosphate (PLP) constitute a ubiquitous class of
biocatalysts, taking part in vital processes of cellular metabolism [Singh & Banerjee,
2011]. Molecular basis of their functions is intensively studied, however the majority of
researches were carried out with E. coli enzymes. Recently functional and structural
characteristics of PLP-dependent enzymes from pathogenic microorganisms became an
object of even higher interest [Astegno et al., 2013, Raj et al., 2013, Revtovich et al., 2012].
The fact that some of these proteins are not presented in mammalian cells makes them a
perspective target for new antibiotics.
O-acetyl-L-homoserine sulfohydrolases are PLP-dependent sulfide-utilizing enzymes in
the L-cysteine and L-methionine biosynthetic pathways of various enteric bacteria and
fungi [Tran et al., 2011]. Up to date there is a lack of structural data on this class of
enzymes - only two 3D structures in RCSB databank.
Here we present for a first time crystal structure of O-acetyl-L-homoserine sulfohydrolase
from bacteria Brucella melitensis (AGSul) - a serious mammalian pathogen, causing
undulant fever. The structure in P3221 space group was refined at 2.0A resolution to
Rf=19.4% and Rfree=24.7%. Despite the asymmetric unit contains two monomers,
detailed contact analysis revealed that protein in crystal is in homotetrameric form which
is in consistent with solution studies. A search for structurally homologous proteins
revealed that AGSul has the same fold as cystathionine γ-lyases. Nevertheless, AGSul
structure possesses some unique features.
This work is supported by grant of Russian Foundation for Basic Research (14-04-31398).
References
Astegno, A. et al. BioMed research international 2013, 701536 (2013)
Raj, I. et al. Biochimica et biophysica acta 1830, 4573-4583 (2013)
Revtovich S.V. et al. Doklady. Biochemistry and biophysics 445, 187-193 (2012)
Singh, S. & Banerjee, R.. Biochimica et biophysica acta 1814, 1518-1527 (2011)
Tran, T. H. et al. Acta crystallographica. Section D, Biological crystallography 67, 831-838
(2011)
95
Poster P-21
Abstract
Serine/threonine-kinases in the malaria parasite Plasmodium
falciparum
Lindner J1, Kapis S2, Betzel C2, Wrenger C1
1Unit
for Drug Discovery, Department of Parasitology, Institute of Biomedical Sciences,
University of São Paulo, Brazil
2Laboratory for Structural Biology of Infection and Inflammation at DESY, University of
Hamburg, Germany
Apicomplexan parasites have a significant impact on health of human and livestock, and
chemotherapy remains a problem. The most severe of these parasites is Plasmodium, the
pathogenic agent of malaria Plasmodium falciparum is proliferating within the human red
blood cells. Thereby the parasite depends on the host metabolism in terms of
carbohydrate, cofactor and amino acids (etc.) supply which have to be imported. Uptake
of these metabolites is either facilitated by transporters or by pinocytosis. The latter is
mediated by cytostome formation followed by lysosomal transport towards the digestive
vacuole. The mechanism which triggers this is unknown in the malaria parasite, however
it has been shown in other organisms that phosphorylation of membrane anchored
proteins play a role. In this sense we focussed on protein kinases which were identified in
the plasmodial genome database. 18 were cloned and analysed for their recombinant
expression profile and subsequently applied for their substrate acceptance. The
discovery of proteins involved in this lysosomal vesicle transport is promising target for
the discovery of novel drugs, taking into account that the parasite relies on metabolic
environment its host.
References
Manning G, In: Genomic overview of protein kinases, WormBook, 1-19 Review (2005)
Flotow H and Roach PJ, The Journal of biological chemistry, 266, 3724-3727 (1991)
Kemp BE and Pearson RB, Trends Biochem. Sci. 15, 342-6 (1990)
96
Poster P-22
Abstract
Structure-function relationships of a novel haloalkane
dehalogenase with two halide-binding sites
Chaloupkova R1, Prudnikova T2,3, Rezacova P4,5, Prokop Z1, Koudelakova T1,
Daniel L1, Brezovsky J1, Ikeda-Ohtsubo W6, Sato Y6, Kuty M2,3, Nagata Y6,
Kuta Smatanova I2,3, Damborsky J1
1Loschmidt
Laboratories, Department of Experimental Biology and Research Centre for Toxic
Compounds in the Environment RECETOX, Masaryk University, Brno, Czech Republic; 2Faculty of
Science, University of South Bohemia in Ceske Budejovice, Ceske Budejovice, Czech Republic; 3Institute
of Nanobiology and Structural Biology, Academy of Sciences of the Czech Republic, Nove Hrady, Czech
Republic; 4Institute of Molecular Genetics, Academy of Sciences of the Czech Republic, Prague, Czech
Republic; 5Institute of Organic Chemistry and Biochemistry, Academy of Sciences of the Czech Republic,
Prague, Czech Republic; 6Graduate School of Life Sciences, Tohoku University, Sendai, Japan
Haloalkane dehalogenases (EC 3.8.1.5) are bacterial enzymes cleaving a carbon-halogen bond by a
hydrolytic mechanism in a broad range of halogenated aliphatic compounds [1]. Novel haloalkane
dehalogenase DbeA from Bradyrhizobium elkanii USDA94 was structurally and biochemically
characterized in this study. The crystal structure of DbeA was solved to 2.2 Å resolution and revealed
the presence of two halide-binding sites. The first chloride-binding site was located in the active site in
between two halide-stabilizing residues. The second chloride-binding site was buried deeply in the
protein core and was approximately 10 Å from the first binding site. This second halide-binding site is
unique to DbeA and has not been previously reported in any other crystal structure of this enzyme
family. To elucidate the role of the second halide-binding site in enzyme functionality, a two-point
mutant (I44L+Q102H) lacking this site was constructed, purified and biochemically characterized. Its
comparison with the wild type enzyme revealed that elimination of the second halide-binding site
decreased stability of the enzyme in the presence of chloride salt and decreased its catalytic activity by
an order of magnitude. Moreover, the two-point substitution resulted in a shift of the substratespecificity class, which is the first time this has been demonstrated for the haloalkane dehalogenase
enzyme family. Changes in the catalytic activity of the variant were attributed to deceleration of the
rate-limiting hydrolytic step, mediated by lower basicity of the catalytic histidine [2].
Figure. Overall structure of the haloalkane
dehalogenase DbeA. C
ribbon trace
represents the elements of the protein
secondary structure. Chloride ions are shown
as grey spheres; chloride ion located in the
active site between two halide-stabilizing
residues is labelled as Cl1; the chloride ion
bound in the second halide-binding site is
labelled as Cl2; the catalytic triad residues are
shown as sticks.
References
1.Koudelakova T, et al., Biotechnol. J. 8, 32-45 (2013)
2.Chaloupkova R, et al., Acta Crystallogr. D Biol. Crystallogr. 70, 1884-1897 (2014)
97
Poster P-23
Abstract
Crystallization and structure determination of peridininchlorophyll a-protein complexes from Symbiodinium sp.
Sommerkamp JAJ1, Machelett MM2, Sniady M1, Mayrhofer J1, Jiang J3,
Blankenship RE4, Hofmann E1
1Protein Crystallography Group, Department of Biophysics, Ruhr-Universität Bochum,
Bochum, Germany;
2Centre for Biological Science, University of Southampton, Southampton, United Kingdom;
3Department of Energy, Environmental & Chemical Engineering, Washington University in
St. Louis, St. Louis, USA;
4Department of Chemistry and Biology, Washington University in St. Louis, St. Louis, USA
The genus Symbiodinium represents the most prevalent group of photosynthetic
dinoflagellates living in endosymbiosis with hermatypic corals and other marine
invertebrates. The soluble peridininchlorophyll a-protein (PCP) is only found in
photosynthetic dinoflagellates and is one of the major light harvesting complexes in these
algae. PCP exists in two different forms, the small form of ~ 15 kDa and the large form of ~
33 kDa. Its primary pigment is peridinin, which is unique for dinoflagellates. We
determined the x-ray structures of two small and two large form PCPs from different
organisms at high resolution (1.1 Å, 1.1 Å, 1.45 Å and 1.95 Å, respectively). Initial
crystallization screening (sitting drop vapor diffusion method) was done with our robotic
nano drop setup. For refinement we used the fourcorners method (Hennessy et al., 2009).
Additionally, hanging drop experiments were set up manually in 24-well plates.
Small PCPs are observed as stable dimers of ~ 15 kDa subunits whereas monomeric
~ 33 kDa apoproteins of large PCPs show high sequence identity of the N- and C-terminal
halves and hence resemble the functional dimer of the small forms. However, in contrast
to small PCPs, all known large PCPs show identical trimer interfaces in the crystal
packaging. The existence and possible functional consequences of this have not yet been
sufficiently investigated in vivo.
Our results show that all structures investigated here, including the pigment cluster, are
structurally similar, however the expression of small and large forms differs between the
studied organisms.
References
Hennessy DN, et al., Acta Crystallogr. D Biol. Crystallogr. 65, 1001-1003 (2009)
98
Poster P-24
Abstract
Design of photovoltaic cell based on metalloproteins a new
approach
Serrano-De la Rosa LE1, Robles-Águila MJ2, Perez
Juárez-Santiesteban H1, Silva-González R3, Moreno A2
KS3,
Stojanoff
V4,
IC-CIDS Benemérita Universidad Autónoma de Puebla, Ed. 130 C, Col. San Manuel, C.P.
72570 Puebla, Pue.,México
2 Instituto de Química, Universidad Nacional Autónoma de México, Circuito Exterior, C.U.
México, D.F. 04510, México.
3 Instituto de Física, Universidad Autónoma de Puebla, Apdo. Postal J-48, Puebla, Pue. C.P.
72570, México
4 Brookhaven National Laboratory, National Synchrotron Light Source, Upton, NY 11973,
USA
1
On this contribution, the Cytochrome C and Azurin proteins were immobilized onto
porous silicon surface using the self assembly technique. The heterostructures were
maintained at ambient conditions through several days. The experimental results show
long term stability of proteins in solid state working as electron-transfer devices. The
atomic force microscopy showed the roughness of the surface similar to both protein
heterostructures (14.5 nm an 11.3 nm respectively) with globular morphology. Analysis of
samples using SEM, showed a porous about 20-24 nm, whereas the cross-section a
thickness between 3.6-3.8 µm. The room temperature fluorescence peak, corresponding
to blue emission, is observed at 362-550 nm. This was due to the silicon through quantum
confinement effect. Raman measurement showed one Raman´s peak; which confirms that
the prepared sample retains the crystallinity of bulk silicon; meanwhile the
immobilization of proteins produce the loss the crystallinity. In reflection spectra, the
porous silicon and changed on refractive index profile at interface porous silicon and
modified surface are shown. The plausible application for developing a photovoltaic cell
using proteins is also presented.
Keywords: Cytochrome –C, Azurin, Porous silicon, Immobilization.
References
Flores-(ernández E et al., Journal of Applied Crystallography 46, 832-834 (2013)
Cruz-Campa JL et al., Sol. Energy Mater. Sol. Cells 95, 551(2011)
Gupta V, Zubia D, Nielson G, Sol. Energy Mater. Sol. Cells 107, 87(2012)
99
Poster P-25
Abstract
Structural and mechanistic insights into the iron processing by
vertebrate ferritins
Pozzi C1, Di Pisa F1, Lalli D2, Bernacchioni C2, Ghini V2, Turano P2,3, Mangani S1,3
1Dipartimento
di Biotecnologie, Chimica e Farmacia, Università di Siena, Siena, Italia 2Dipartimento di
Chimica, Università di Firenze, Firenze, Italia 3Centro Risonanze Magnetiche (CERM), Università di Firenze,
Firenze, Italia.
Ferritins are intracellular proteins that concentrate thousands of iron(III) ions as solid mineral [1]. The
ferritin molecule is a multimeric system forming an external cage around the storage cavity. Iron(II) enters
the protein shell through ion channels, proceeds inside each subunit, reaches the active site, where it is
oxidized by O2 and eventually enters the cavity [2]. In vertebrate ferritins, like the frog M ferritin here
studied, this path is about 40 Å long. Data on iron-ferritin are available for bacterioferritins [3], for the
anaerobe Pyrococcus furious [4] and for the marine pennate diatom Pseudo nitzschia multiseries [5]. Besides
bacterioferritins, where iron can be cofactor and/or substrate, studying iron in ferritins is difficult because
the protein provides only weak interactions to iron, that moves without resting at any tight binding site. The
transient nature of protein-iron interactions translates into undetectable iron binding in the crystals. In
bacterioferritins iron is well detected but its function is controversial [3]. Understanding the protein
capability i) to attract and guide iron along the path, ii) to keep it at the catalytic oxidation site for the
oxidation/coupling reaction time, and iii) to avoid any definitive sequestration of the metal at protein
binding sites is a scientific challenge. A soaking/flash freezing method has been developed to allow aerobic
and anaerobic addition of iron(II) to frog [6] and human ferritin crystals (figure 1). Multi-wavelength
anomalous diffraction data have been exploited to unambiguously detect the iron atoms. The method has
allowed us to observe for the first time the iron binding sites of a vertebrate ferritin and to see how they
evolve with time.
Moreover, we have studied the iron uptake process by collecting X-ray crystallographic data at room
temperature. Iron loading has been performed on protein crystals by free-metal diffusion in capillary and
the development of the catalytic reaction has been monitored. The structural data, together with stop-flow
kinetic data, provide new clues about the iron processing mechanism by ferritins.
Figure 1. Iron (orange spheres) bound to the ferroxidase site (A) and additional sites (B) in human ferritin.
References
[1] Liu X, Theil EC, Acc. Chem. Res. 38, 167-175 (2005)
[2] Tosha T, Ng HL, Bhattasali O, et al., J Am Chem Soc. 132, 14562-14569 (2010)
[3] Le Brun NE, Crow A, Murphy MEP, et al., Biochim. Biophys. Acta, 1800, 732-744 (2010)
[4] Tatur J, Hagen WR, Matias PM. J Biol Inorg Chem, 12, 615-630 (2007)
[5] Marchetti A, ParkerMS, Moccia LP, et al,. Nature, 457, 467-470 (2009)
[6] Bertini I, Lalli D, Mangani S, et al., J Am Chem Soc, 134, 6169-6176 (2012)
100
Poster P-26
Abstract
Crystallization and structural characterization of haloalkane
dehalogenase LinB mutants
Degtjarik O1,3*, Iermak I1*, Chaloupkova R5, Rezacova P4, Kuty M2, Damborsky J5 and
Kuta Smatanova I1,3
1University
of South Bohemia, Faculty of Science, Ceske Budejovice, Czech Republic;
of South Bohemia, Faculty of Fisheries and Protection of Waters, Institute of
Complex Systems and CENAKVA, Nove Hrady, Czech Republic; 3Academy of Sciences of the
Czech Republic, Institute of Nanobiology and Structural Biology GCRC, Nove Hrady, Czech
Republic; 4Institute of Molecular Genetics of the Academy of Science of the Czech Republic,
16637 Prague, Czech Republic; 5Loschmidt Laboratories, Faculty of Science, Masaryk
University, Brno, Czech Republic
* equal contribution
2University
LinB is a microbial enzyme of the haloalkane dehalogenase family that catalyse cleavage of
the carbon-halogen bond in halogenated aliphatic pollutants, resulting in the formation of
a corresponding alcohol, a halide ion and a proton. Haloalkane dehalogenase LinB isolated
from a bacterium Sphingobium japonicum UT26 has relatively broad substrate specificity
and can be potentially used for biosensing and biodegradation of environmental
pollutants. Different variants of haloalkane dehalogenase LinB were constructed with a
goal to study the effect of mutations on the enzyme functions. Mutations in LinB73
(D147C+L177C) cause blocking of the main tunnel by formation of disulphide bridge. In
LinB86 (140A+143L+177W+211L) variant mutations lead to blocking of the main tunnel
and opening the alternative way for connection the deeply buried active site with the
surrounding solvent.
Both LinB variants were successfully crystallized and full data sets were collected. Crystals
of LinB86 were obtained in Index (Hampton Research, USA) and Morpheus (Molecular
Dimensions Ltd., UK) screens. LinB86 crystals diffract to resolution of 2.34 Å and belong to
H-centered trigonal H32 space group. Crystals of LinB73 were found in PEGs Suite
(QIAGEN, Netherlands) and data set was collected up to resolution 1.15 Å. They belong to
P212121 space group. Structures of both mutants were solved by molecular replacement
using the coordinates from LinB32 (L177W) variant and LinB wild type. The structure
analysis of both variants will follow.
This research was supported by the GA CR (P207/12/0775), GAJU (141/2013/P) and ME
CR (CZ.1.05/2.1.00/01.0024).
101
Poster P-27
Abstract
Crystallization of mutants of brazzein, a sweet protein
Tae-Yeon K1,2, Tae-Sung Y1,2
1Bio-Analytical
Science, University of Science & Technology, KOREA; 2Medical Proteomics
Research Center, Korea Research Institute of Bioscience & Biotechnology, KOREA
Some proteins are known to stimulate taste receptors and elict sweet taste. Thaumatin,
monellin, mabinlin, pentadin, curculin, miracullin and brazzein have been reported as
such sweet proteins [1]. Brazzein originates from Pentadiplandra brazzeana and its crystal
structure was recently reported and compared with its NMR structures [2,3]. It has been
known that mutations of brazzein either abolish or increase sweetness [4]. To study
crystal structures of brazzein mutants, we synthesized gene construct based on wild-type
brazzein from NCBI database (KF013250.1). It was cloned in E. coli DH10β strain with
pET-21a vector. Its protein was expressed in Rosetta-gami B(DE3) strain. Brazzein
mutants (D29A, D29K E41K and R43A) were also expressed in Rosetta-gami B(DE3)
strain with site-directed mutagenesis of the recombinant brazzein. The expressed proteins
were purificated by His-tag affinity chromatography, cation exchange and gel filteration
chromatography. Among them, R43A mutant resulted in two crystal forms,
a = b = 51.5
Å, c = 60.6 Å, = = 90°, γ=120° in P3121 and a = 51,3 Å, b = 89.1 Å,
c = 121.2 Å,
= = γ = 90° in P212121 from the same crystallization condition (1.0 M Citrate buffer,
pH=4, 1.0 M NaCl).
References
[1] Kant R, Nutrition Journal, 4:5(2005)
[2] Sannali M. Dittli et al., Chem. Senses, 36, 821-830 (2011)
[3] Nagata K et al., Acta Cryst. D, Apr, 69, 642-7 (2013)
[3] Hellekant G and Danilova V, Chem.Senses, 30, i88-i89 (2005)
102
Poster P-28
Abstract
Batch crystallisation of model proteins with 3D nanotemplates
Chum KKCƚ, Chang WCƚ, Amberg Cƚ, Shah UVƚ, Ho TCTǂ, Ricard Fǂ, Williams DRƚ,
Heng JYYƚ
ƚSurface and Particle Engineering Laboratory, Department of Chemical Engineering, Imperial
College London, London, U. K
ǂParticle Sciences and Engineering UK, GlaxoSmithKline R&D, Stevenage, U. K
Heterogeneous seeding has been employed as a technique in increasing the likelihood of obtaining
protein crystals. 3D nanotemplates, mesoporous silica of specifically designed surface properties
(pore size, surface roughness and surface chemistry), have been demonstrated to be effective in
promoting protein crystallisation in vapour diffusion. Shah et al (2012) proposed and
demonstrated that nanotemplates of pore diameter close to the size of protein are most efficient in
promoting protein crystallisation. Results to date demonstrate the applicability of this concept to
the batch crystallisation of two model proteins, hen egg-white lysozyme and bovine liver catalase.
At a working volume of 400 mL, the addition of nanotemplates with 3.5 ± 1.0 nm (0.1 g) yielded a
crystallisation onset 0.08 times (20 minutes) that of unseeded crystallisation experiments (255
minutes) for lysozyme (dimensions = 3.0 x 3.0 x 4.5 nm). The use of nanotemplates with larger
pore diameters resulted in a crystallisation onset 0.22-0.35 times the unseeded experiments.
Extending beyond lysozyme, non-agitated batch crystallisation (2 mL) was performed on catalase;
a model protein of hydrodynamic diameter 90 Å. Nanotemplates of pore diameters 11.0 ± 3.0 nm
was demonstrated to be superior heterogeneous seed to those of pore diameters 3.5 ± 1.0 nm at a
range of catalase concentration studied (4 – 10 mg/mL). This effect is more prominent in lower
supersaturation levels.
Above studies on model proteins demonstrates the importance of specific pore design in reducing
the nucleation time for the batch crystallisation of proteins. This method would be applicable to
the biopharmaceutical industries, where the newly discovered molecules are not expressed in
high concentrations.
References
1.U. V. Shah et al, Cryst. Growth. Des. 12, 1362-1369 (2012)
2.U. V. Shah et al, Cryst. Growth Des. 12, 1772-1777 (2012)
103
Poster P-29
Abstract
Protein crystallization in a new meso oscillatory flow reactor
Castro F1, Ferreira A1,2, Rocha F2 and Teixeira JA1
1Centre
of Biological Engineering, Dep. of Biological Engineering, Univ. of Minho, Braga, Portugal; 2Faculty of
Engineering of Porto, Dep. of Chemical Engineering, Univ. of Porto, Porto, Portugal
Protein crystallization is best known for its use in protein three-dimensional (3D) structure determination by X-ray
diffraction. Little systematic know-how exists for protein crystallization as a process step (Schmidt et al.). However,
protein crystallization offers great potential to be used for separation and formulation applications on an industrial
scale (Schmidt et al., Parambil et al.). Purification of proteins via crystallization may offer significant cost reductions
compared to conventional downstream processing techniques such as chromatography. Moreover, protein crystals have
a longer storage life and greater purity compared to the dissolved form.
Protein crystallization as a process step requires rapid and quantitative crystallization. However, most of the studies in
protein crystallization had been focused on producing high quality crystals obtained by slow crystallization from small
volumes under static conditions. But, crystallization under static conditions is often diffusion-limited, leading to rather
long process durations and/or low yields. Hence, protein crystallization under flow conditions has attracted attention
with the capability to improve the convective protein transport, reducing the crystallization time and improving yield
(Parambil et al.).
In this context, micro- (microliters) and meso- (milliliters) reactors systems offer unequalled experimental conditions to
explore the complexity of protein crystallization. Indeed, scaling down offers several advantages, namely enhanced heat
and mass transfer, better control over the contact mode of the reagents and optical access for in situ characterization
(Leng & Salmon). As only small amounts of protein are often available, experimentation on the liter scale is ruled out,
whereby representative experiments in scale-down reactors can provide important information for industrial-scale
crystallization (Schmidt et al.).
The purpose of the present work is to apply protein crystallization as an alternative purification step for proteins of
pharmaceutical or industrial interest. For this, we propose to investigate protein crystallization under flow conditions,
more precisely in a new meso oscillatory flow reactor (OFR) based on Reis work (Reis). The reactor consists of a glass
jacketed tube provided with smooth periodic cavities (SPC) and is operated under oscillatory flow mixing, controlled by
the oscillation frequency and amplitude.
Figure 1: Geometry of the SPC tube composing the
meso-OFR (Reis)
First experiments were focused on the study of favourable crystallization conditions for a model protein, lysozyme.
Lysozyme from egg white and precipitating agent (sodium chloride) solutions were both prepared in sodium acetate
buffer at pH 4.0. Both solutions were filtered (0.22 µm) and tempered at 20 ºC. Crystallization assays were started by the
simultaneous injection, through a syringe pump, of lysozyme and sodium chloride solutions in the meso reactor, which
was temperature controlled (T = 20 ºC). Through light transmittance readings and optical microscopy images, it was
possible to determine induction times and crystallization times, as well as shape and size of the crystals.
References
Schmidt S, et al., Engineering in Life Sciences 5, 273–276 (2005)
Parambil JV, et al., Crystal Growth & Design 11, 4353–4359 (2011)
Leng J & Salmon JB, Lab on a chip 9, 24-34 (2008)
Reis N, PhD Thesis, Braga, Portugal (2006)
104
Poster P-30
Abstract
Advantages of under-oil crystallization
Kulathila R, Xie X
Novartis Institute for Biomedical Research, Cambridge, Massachusetts, USA
Using oil in crystallization and in cryo-protection has advantages (1, 2). Recent advances
in automation allow minimal amount of protein with a maximum number of
crystallization conditions to be screened in a high-throughput robot such as the Mosquito
ttplabtech robot and others. Under-oil crystallization reduces vaporization of the proteinmother liquor drop mix during the high-throughput automation setup. Furthermore, oil
can be used as a filling agent to reduce the protein volume in a high-throughput capillary
tray such as the Crystal Former tray (Microlytic). The benefit of using oil is also
demonstrated in capillary crystallization. Using oil to replace the seal at the top end of the
capillary allows counter diffusion at varying rates (1,3). A variety of oils, owing to their
different vaporization rates, enable new crystallization kinetics. In certain oils, one can
even scoop the crystals directly from the under-oil tray and cryo-preserve them without
the tedious process of searching for a cryo buffer. The following crystallization
experiments demonstrate the success of using oil in the crystallization setups of three
proteins complexed with inhibitors. In particular, oil vaporization coupled with counter
diffusion allowed us to obtain diffracting crystals of the proteins with and without
inhibitors, which could otherwise not have been obtained.
References
1)Hampton Research, http://hamptonresearch.com/documents/growth_101/6.pdf
2)Chayen NE, et al, Journal of Crystal Growth 122 (1992) 176-180
3)J.A. Gavira D, et al,
http://www.chemie.uni-hamburg.de/bc/betzel/Gavira-Gallardo_1.pdf
105
Poster P-31
Abstract
Microcrystallization of the fluorescent protein IrisFP in view of
time-resolved XFEL serial femtosecond crystallography
Guillon V1, Colletier J-P1, Schiro G1, Woodhouse J1, Byrdin J1, Adam V1, Bourgeois D1,
Weik M1
1Institut de Biologie Structurale, Univ. Grenoble Alpes – CNRS - CEA , F-38044 Grenoble,
France
Phototransformable fluorescent proteins (PTFPs) are invaluable tools in advanced
fluorescence microscopy of live cells (including single-molecule localization microscopy)
because their fluorescence emission intensity and/or color can be controlled by
irradiation with light of a specific wavelength. IrisFP is a PTFP that can be photoconverted
from green to red. In both forms, it can be switched reversibly between an on- and an offstate [1]. This bi-photochromism makes IrisFP a most versatile candidate for two-color
pulse-chase single-molecule localization microscopy [2]. Photoswitching involves
chromophore isomerization, proton transfer and rearrangements of side chains in the
chromophore pocket. The static structures of IrisFP in its on- and off-states have been
resolved by synchrotron X-ray crystallography [1], but the structures of photoswitching
intermediates remain elusive. Only time-resolved serial femtosecond crystallography
(SFX) [3, 4] can provide structural insights into such short-lived intermediates states.
Owing to the technical requirements of the envisioned experiment and to the decreased
penetration depth of visible light within protein crystals, a prerequisite is to produce large
amounts of microcrystals. The initial crystallization protocol yielding macrocrystals [1]
has been modified so as to generate myriads of smaller crystals (10 × 10 × 50 – 90 mm).
The new crystals diffract to high resolution, as evidenced by recent data collection at the
ESRF synchrotron (1.7 Å). In crystallo spectroscopic experiments demonstrate that the
new crystals are photoswitchable, with a green-off state half-life similar to that
determined earlier on macrocrystals, which indicates that they are amenable to the
proposed time-resolved SFX experiments.
This piece of work is part of a larger ongoing project involving many collaborators,
including Michel Sliwa, Marco Cammarata, Ilme Schlichting, Thomas Barends, Robert
Shoeman, Bruce Doak, Karol Nass, Sabine Botha, Lutz Foucar, Leonard Chavas, FrançoisXavier Gallat, Mishiro Sugawara, and Sebastien Boutet.
[1] Adam V, et al. (2008) Proc Natl Acad Sci U S A 105: 18343-18348
[2] Fuchs J, et al. (2010) Nature Methods 7, 627-630
[3] Chapman HN, et al. (2011) Nature 470: 73-77.
[4] Aquila A, et al. (2012) Optics express 20: 2706-2716
106
Poster P-32
Abstract
Establishment of a protocol for robust production and
crystallisation of mineralocorticoid receptor ligand binding domain
Bäckström S1, Aagaard A1, Edman K1
1Discovery Sciences, AstraZeneca R&D, Mölndal, Sweden
The mineralocorticoid receptor (MR) is an attractive drug target for the treatment of a
number of diseases such as hypertension and heart failure. It has been shown that soluble
expression of the MR ligand binding domain (LBD) in E. coli can be accomplished in
presence of high affinity agonistic ligands. Furthermore, a point mutation (S810L) also
makes expression of MR-LBD in complex with antagonistic ligands possible (Bledsoe et al.,
2005). We have combined these techniques with the C-terminal tethering of a peptide
from a co-regulator protein. This approach resulted in a dramatic increase in the protein
expression enabling higher throughput of MR-ligand complex structure determination.
The availability of large amounts of protein allowed for an extensive crystallisation
screening campaign resulting in crystals from one unique crystallisation condition.
References
Bledsoe R.K. et al., J .Biol. Chem 280, 31283-31293 (2005)
107
Poster P-33
Abstract
Application of the silkworm expression system for the structural
biology: A case study of human CD1d-b2m
Kita S1, Kusaka H1, Yoshida K1, Kasai Y1, Niiyama M2, Sugiyama S2, Murata M2, Kuroki
K1, Maenaka K1
1Pharmaceutical
2Graduate
Sciences, Hokkaido University, Hokkaido, Japan
School of Engineering, Osaka University, Osaka, Japan
Producing recombinant proteins in native conformation and function is one of the most important parts in
structural biology. Recently, demands for the structure analysis of mammalian proteins, especially for
membrane proteins and multifactor complexes are gradually increased. Baculovirus-insect cell expression
system is one of the widely used expression system to meet such demands. Up to now, insect cell lines such
as Sf9, Sf21, S2 and Hi5 cells are generally used in baculovirus-insect cell expression system. Meanwhile, we
previously developed the Baculovirus-silkworm expression system and succeeded in the expression of large
amounts of membrane proteins. The silkworm expression is easy to handle, less contamination and
comparatively low-cost method. We established this method by modifying the Bac-to-Bac expression system
(Invitrogen) (Figure 1). In the system, plasmids harboring gene of interest are introduced into E. coli host
BmDH10Bac. In BmDH10Bac cells, gene of interest is transpositioned from the plasmid to a baculovirus shuttle
vector, bacmid. Recombinant bacmids are isolated from BmDH10Bac cells and are subseqentlly injected to
silkworms, Bombyx mori. Recombinant proteins and viruses are produced in several days. Here we represent the
application of this system to the preparation and structure analysis of human CD1d-β2m heterodimeric
complex. CD1d belongs to the non-classical major histocompatibility complex (MHC) class I protein and
present the various types of glycolipids to the natural killer T cells and T cells. To elucidate the molecular
basis for the CD1d protein, we prepared human CD1d protein using silkworm expression system and solved
its crystal structure.
The extracellular domains of CD1d and β2m are subcloned into a plasmid, pFastBac Dual. BmDH10Bac
competent cells were transformed by the pFastBac Dual plasmids and recombinant bacmids were obtained.
Recombinant bacmids were injected to silkworm, Bombyx mori. The hemolymph was collected after 6 days
and purified by ammonium sulfate precipitation followed by Ni-affinity chromatography and size exclusion
chromatography. After purification, the homogeneity of purified protein was clarified by SDS-PAGE to be
>90%. Finally 1 mg of proteins was obtained from 50 silkworm larvae. The screening of crystallization
conditions was performed with JCSG Core suites kit (QIAGEN) by the sitting-drop vapor-diffusion method
using mosquito robot (TTP labtech). Crystals of CD1d-β2m were obtained in a condition containing
ammonium sulfate as precipitant. The crystals were diffracted to 4.15 Å and X-ray diffraction dataset was
collected on the beamline BL-5A of Photon Factory (Tsukuba, Japan). The crystals belong to spacegroup
I213, a = b = c = 240.4 Å. The structure of CD1d-β2m was solved by molecular replacement program phaser
using the structure previously reported (PDB ID: 4LHU) as a search model. The refinement of the model is
now proceeding.
Figure 1. Schematic illustration of the silkworm expression system.
References
Tomoko M, et al., BBRC., 326, 564-569 (2005)
Mizuho K, et al., BBRC., 385, 375-379 (2009)
Sasaki K, et al., BBRC., 387, 575-580 (2009)
108
Poster P-34
Abstract
Crystallization of human hypoxanthine-guanine
phosphoribosyltransferase in the presence of GMP or acyclic
nucleoside phosphonates
Wang TH 1, Keough DT1, Hocková D 2, Guddat LW1.
1The
School of Chemistry and Molecular Biosciences, The University of Queensland, Brisbane
4072, Queensland, Australia
2Institute of Organic Chemistry and Biochemistry, Academy of Sciences of the Czech Republic, v.v.i.
Flemingovo nám. 2, CZ-166 10 Prague 6, Czech Republic
Hypoxanthine-guanine-phosphoribosyltransferase (HGPRT) catalyses the conversion of
hypoxanthine or guanine to inosine monophosphate (IMP) or guanosine monophosphate (GMP).
The presence of a divalent metal ion such as Mg2+ is required for the reaction to occur. HGPRT
plays a crucial role in the synthesis of purine nucleotides via the purine salvage pathway [1,2]. In
many organisms, including Plasmodium parasites (the causative agents of malaria), the activity of
this pathway appears to be essential for their survival. As a result, it is an attractive target for drug
discovery and design. One class of compound that has been developed into potent inhibitors of
this enzyme is the acyclic nucleoside phosphonates (ANPs), which are analogs of the products of
the reaction [3].
In general, the crystallization of the parasite enzymes have been challenging whereas the human
enzyme, while although also difficult to crystallize, has been more amenable in this endeavour. For
that reason, the focus of the current study is on the human enzyme. There are several reasons that
make crystallization of this enzyme difficult. They include the fact that (i) human HGPRT is a
tetramer and contains several free thiol groups; (ii) there are large sections of the polypeptide that
are inherently flexible; and (iii) the ANPs that have been designed as inhibitors and they may need
to also coordinate with one or two Mg2+ ions for optimal binding. To optimize expression and
purification of the protein, the plasmid containing the cDNA coding for this enzyme was produced
(by GeneART) so as to include a hexa-histidine tag at the amino terminus. Three of the four
cysteine residues (C22, C105 and C205) were mutated to alanine to prevent inappropriate
disulfide bond formation and to increase stability. The fourth, C65, was unchanged as mutation of
this residue resulted in inactive enzyme [3]. Using this construct, up to 100 mg of purified enzyme
could be produced from eight litres of culture. For crystallization trials, the Mosquito robot was
used to set-up the trays and the experiments were housed in the FORMULATRIX incubator and
imaged with the Rockimager software. Once initial crystals were obtained, optimization was
performed on a larger scale using VDX plates. Even though the structures of the inhibitors varied
only slightly, in most cases new crystallization conditions were required for each enzyme-inhibitor
complex. Analysed together the results make for an interesting case study.
References
[1][Reyes P, et al., Mol Biochem Parasitol 5, 275-290(1982)
[2],Gardener M, et al., Nature 419, 498-511 (2002)
[3]Keough DT, et al., J Med Chem 52, 4391-4399(2009)
109
Poster P-35
Abstract
Precise protein crystallography with the Echo® 550 liquid handler
Zahr D1, Edwards B, Harris D, Datwani S and Barco J
1Labcyte Inc., USA
Acoustic transfer with the Echo liquid handler simplifies the small scale crystallography
process by transferring reagents precisely at 2.5 nL increments. Using Dynamic Fluid
Analysis™, the Echo liquid handler can adjust transfer settings on the fly on a well-by-well
basis. This allows a single fluid class setting to be utilized for a wide variety of viscous and
osmotic fluids, greatly simplifying the liquid transfer process. We demonstrate an
expanded ability to transfer a wide variety of reagents from various protein
crystallography sparse matrix screens. To enable the potential for on the fly grid screens,
we demonstrate the precise transfer of viscous reagents (including high molecular weight
PEGs) and organic solvents (including MPD).
50 nL (Echo 555)
100 nL(Echo555)
1 µL (manual)
Protein crystal hits generated in 96-well Swissci microplates by adding Hampton Crystal
Screen HT reagents to protein solutions on the Echo 555 liquid handler and by hand. Top
row: 50 mg/mL lysozyme solution + E1 reagent (2.0 M sodium chloride, 10% w/v
Polyethylene glycol 6,000). Bottom row: 25 mg/mL thaumatin solution + C5 reagent
(0.1 M HEPES sodium pH 7.5, 0.8 M Potassium sodium tartrate tetrahydrate).
110
Poster P-36
Abstract
Crystal structures of the PH domain from p190-RhoGEF bound to
activated Rho or Rac GTPase suggest a novel cross-talk regulation
between Rho and Rac signalling pathways
Chen Z1, Dada O2, Gutowski S2, and Sternweis PC2
1Department
of Biophysics, 2Department of Pharmacology, University of Texas, Southwestern
Medical Center, Dallas, Texas, USA
The Rho family of monomeric GTPases belongs to the Ras superfamily and is composed of several
proteins including Rho, Rac and Cdc42. They control a wide range of cellular processes, and cycle
between an inactive GDP-bound state and an active GTP-bound state. This cycling is tightly
regulated by sets of protein partners. Activation of Rho largely depends on Rho Guanine
nucleotide Exchange Factors (RhoGEFs), which catalyze the exchange of GTP for GDP on Rho. The
exchange activity resides within the tandemly linked Dbl homology (DH) and plekstrin homology
(PH) domains of these RhoGEFs. Recently, the PH domains from the Lbc-family of RhoGEFs (LbcRhoGEFs) were shown to interact directly with activated Rho bound to GTP1,2. This was
unexpected and has been proposed to function as a positive feed back for Rho signaling. All three
family members of Rho GTPases, RhoA, RhoB and RhoC, interact directly with the PH domains
from Lbc-RhoGEFs. One member of the Lbc-RhoGEFs, p190-RhoGEF, also binds to activated Rac1
via its PH domain, albeit with much lower affinities. We have crystallized and solved structures of
PH domain from p190-RhoGEF bound to RhoA-GTP, or Rac1-GTP. The interaface between
activated Rac1 and the PH domain is analagous to the one between activated RhoA and PH,
utilizing the same hydrophobic surface on the PH domain. Similar to RhoA, activated Rac1
interacts with the PH domain via its effector-binding surface. However, the Rac1-PH interface
buries less surface area than the interface between RhoA and PH. Activated Rac1 can stimulate the
RhoGEF activity of p190-RhoGEF in vitro, and mutations of key hydrophobic residues in the PH
domain abolish this activation. These structures, together with recent structures of PH:RhoA
complexes, identify a highly conserved interface between PH domains from Lbc-RhoGEFs and
activated small GTPases. More importanly, the Rac1:PH interaction reveals a potential mechanism
for novel cross-talk regulation between the Rho and the Rac signaling pathways.
Ribbon diagram depicting the structure of the
complex of p190RhoGEF-PH domain (light
grey) bound to activated Rac1 (dark grey). GTP
and magnesium bound to Rac1 are shown as
ball and stick models.
References
1.
Chen, Z, et al., J. Biol Chem 287: 16369-77 (2012)
2.
Medina, F, et al., J. Biol Chem 288: 11325-33 (2013)
111
Poster P-37
Abstract
SONICC results from the field: How nonlinear optical microscopy
has revolutionized crystallization screening and aided in
microcrystalline detection.
Fisher M1, Gualtieri EG1
1Formulatrix, Inc, Waltham, MA, USA
Crystallization screening processes can often be a time consuming and frustrating process. When
viewing images of crystallization plates, it is not often apparent if protein crystals have formed,
especially if they are very small or obscured by a turbid matrix. SONICC (Second Order Nonlinear
Imaging of Chiral Crystals) was developed to facilitate the detection of microcrystals even when
buried in precipitate and to differentiate between salt crystals and protein crystals (Figure 1).1-4
SONICC is now used routinely at over 20 pharmaceutical and academic laboratories worldwide
finding crystallization hits that would never be investigated with conventional techniques. This
talk will briefly discuss the nonlinear optical technology utilized in SONICC and then focus on
results in the field. We will summarize how SONICC has impacted the community over the past
few years and how some labs now heavily rely on SONICC and have completely changed their
workflow in crystallization screening.
Figure 1. A LCP drop imaged with conventional techniques (visible, crossed polarizers, UV) and
SONICC (UV-TPEF and SHG). The SHG imaging mode identifies microcrystals that could
not be identified with other imaging modalities.
References
1.
Haupert, L.; Simpson, G. J. Methods 55, 379-386 (2011)
2.
Kissick, D. J, et al., Ann. Rev. Anal. Chem. 4, 419-37 (2011)
3.
Madden, J. T, et al., Acta Crystallographica D D67, 839-846 (2011)
4.
Kissick, D. J, et al., Anal. Chem. 82(2), 491-497 (2011)
112
Poster P-38
Abstract
Structure determination of enzymes involved in mutagenesis in
Mycobacterium tuberculosis
Broadley SG1,2, Warner DF2,3 Sewell BT1,3
1Structural
Biology Research Unit, University of Cape Town, South Africa; 2 MRC/NHLS/UCT Molecular
Mycobacteriology Research Unit and DST/NRF Centre of Excellence for Biomedical TB Research, University
of Cape Town, South Africa; 3Institute for Infectious Disease and Molecular Medicine, University of Cape
Town, South Africa
Drug-resistant strains of Mycobacterium tuberculosis (MTB) threaten a global tuberculosis (TB) control
effort that is founded on chemotherapeutic intervention in active, infectious disease. All drug resistance in
MTB arises through the acquisition and maintenance of spontaneous mutations in antibiotic target or
related genes. This observation places enormous importance on the need to understand the mutagenic
mechanisms operating in MTB and, in turn, identifies mutagenic pathways as a compelling target for novel
anti-TB drugs. Previous work has implicated the DNA damage-inducible C family DNA polymerase, ImuC
(also known as DnaE2), in virulence and the emergence of antibiotic-resistant MTB mutants in vivo. ImuC
operates as part of a three-component “mutagenic cassette” comprising the essential accessory proteins,
ImuB (Rv3394c) – a pseudo Y-family polymerase – and ImuA’ (Rv3395c), a RecA-like protein of unknown
function.
The aim of this study is to obtain crystal structures of all three proteins in order to gain insight into the
geometry of the ImuC active site and to elucidate potential interacting domains in ImuC, ImuA’, and ImuB.
To date, ImuA’ has been successfully expressed in E. coli with a maltose binding protein tag, purified, and
placed into crystallization trials. A possible crystal hit has been obtained which is currently in the process of
optimization.
Given previous evidence from yeast two hybrid assays that ImuA’ and ImuC interact with the extended Cterminal region of ImuB, additional protein expression systems have been selected to test the potential for
co-expression of the mutagenic cassette proteins in different combinations.
Once soluble protein is obtained, the proteins will be placed into crystal trials separately however cocrystallisation of the interacting proteins will be attempted as well as co-crystallization of ImuC with DNA.
DNA polymerases bind double stranded DNA but have little sequence preference. In order to capture ImuC
together with DNA it is necessary to design a DNA primer and template that would result in a doublestranded region with a 5’ overhanging single stranded portion. Catalysis will need to be prevented without
mutating essential residues in the polymerase as it is important for our study to investigate the active site
intact.
Crystallization conditions from Hampton will be screened using a mosquito pipetting robot. Any crystal hits
will be identified visually and optimised by manual set up of hanging drop vapour diffusion. Optimisation
will be performed by screening pH and precipitant concentration in a 24 well plate.
References
Liu B, et al., Structure 21, 658-664 (2013)
McHenry CS, EMBO Rep. 12, 408–414 (2011)
Warner DF, et al., Proc. Natl. Acad. Sci. U S A 107, 13093–13098 (2010)
Wing RA, et al., J. Mol. Biol. 382, 589-869 (2008)
Galhardo RS, et al., Crit. Rev. Biochem. Mol. Biol., 42, 399-435 (2007)
Boshoff HIM, et al., Cell 113, 183–193 (2003)
113
Poster P-39
Abstract
Improvement of high throughput crystallization screening system
at KEK-PF in Japan
Kato R1, Hiraki M2, Yamada Y1, Kawasaki M1, Matsugaki N1
1Structural
2Mechanical
Biology Research Center, Photon Factory, IMSS, KEK, Tsukuba, Japan;
Engineering Center, Applied Research Laboratory, KEK, Tsukuba, Japan
Search for crystallization conditions is still bottleneck for the X-ray crystal structure
analysis. To overcome this, we developed a high throughput crystallization system and
have been operating it. In the PDIS (Platform for Drug Discovery, Informatics, and
Structural Life Science) project in Japan, we receive any protein samples from PDIS users
and subject them to the crystallization system.
The PDIS project is divided into three parts; “Structural Analysis”, “Drug Discovery” and
“Information.” “Structural Analysis” consists of four areas; “Functional Genomics”, “Protein
Production”, “Structural Analysis” and “Bioinformatics.” Each area is composed of several
universities and institutes in Japan. Our crystallization system belongs to “Protein
Production” area and carries out crystallization screening. Protein crystallography
beamlines at KEK-PF belong to “Structural Analysis” area, and both of the crystallization
system and the beamlines form a pipeline in a unified fashion for structural life science.
We are improving our crystallization screening system by installing a new crystal
observation method by SONICC (Second Order Non-linear Imaging of Chiral Crystals), a
small volume dispensing system, and so on. These advanced technologies will support
users in the pipeline. In addition, in strong conjunction with protein crystal production
and beamline activity in the same campus, some collaborations are scheduled to be
developed, and they will contribute the efficiency of the pipeline.
References
Hiraki M, et al., J Synchrotron Radiat. 20, 890-893 (2013)
Yamada Y, et al., J Synchrotron Radiat. 20, 938-942 (2013)
Chavas L, et al., J Synchrotron Radiat. 19, 450-454 (2012)
Hiraki M, et al., Acta Crystallogr D 62, 1058-1065 (2006)
114
Poster P-40
Abstract
Structural basis of exopeptidase activity of a dipeptidyl
aminopeptidase BII, a member of the family S46 peptidases
Sakamoto Y1,7, Suzuki Y2,7, Iizuka I1, Tateoka C1, Roppongi S1, Fujimoto M1, Inaka K3,
Tanaka H4, Masaki M5, Ohta K5, Okada H2, Nonaka T1, Morikawa Y2, Nakamura KT6,
Ogasawara W2, Tanaka N6
1School
of Pharmacy, Iwate Medical University, Yahaba, Japan; 2Department of Bioengineering, Nagaoka
University of Technology, Nagaoka, Japan; 3Maruwa Foods and Biosciences Inc., Yamatokoriyama, Japan;
4Confocal Science Inc., Tokyo, Japan; 5Japan Aerospace Exploration Agency, Tsukuba, Japan; 6School of
Pharmacy, Showa University, Tokyo, Japan; 7Contributed equally to this work
The dipeptidyl aminopeptidase BII (DAP BII) from Pseudoxanthomonas mexicana WO24, originally isolated
from the waste of a tofu factory (Ogasawara et al., 1996), belongs to a new class of serine peptidases, family
S46 (Suzuki et al., 2014). The amino acid sequence of the catalytic unit of DAP BII exhibits significant
similarity to those of clan PA endopeptidases, such as chymotrypsin, which form the largest group of known
peptidases. However, the molecular mechanism of the exopeptidase activity of DAP BII is unknown. We
have determined crystal structures of DAP BII in its peptide-free form and in seven peptide-bound forms
(Sakamoto et al., 2014). For peptide-free form, higher resolution data were obtained from a space-grown
crystal (1.95 Å resolution) than from a ground-grown crystal (2.20 Å resolution). The space-grown peptidefree crystals were obtained using a counter-diffusion method (Garcia-Ruiz & Morena, 1994) under a
microgravity environment in the Japanese Experimental Module “Kibo” at the International Space Station
(Takahashi et al., 2013).
DAP BII contains a peptidase domain including a double b-barrel fold, which is characteristic of the
chymotrypsin superfamily, and an unusual, previously unreported a-helical domain (Figure 1). This is the
first structural example of an exopeptidase containing both a catalytic chymotrypsin fold and a regulatory
domain. Structures of a peptide complex revealed the following two important functions of the a-helical
domain: (i) the domain covers the active-site cleft where the catalytic triad (His86, Asp224, and Ser657) is
located and limits the access of globular substrates to the active site, and (ii) the side chain of Asn330 in the
domain forms a bifurcated hydrogen bond with the main chain of the N-terminal residue of the bound
peptide. The side chains of Asn215, Trp216, and Asp674 in the catalytic domain are also involved in the
recognition of the substrate N-terminus. These observations clearly indicate that the a-helical domain
regulates the exopeptidase activity of DAP BII. Moreover, a series of peptide-bound structures provide
insights into the substrate-recognition and catalytic mechanisms of DAP BII. Because S46 peptidases are not
found in mammals, we expect that these enzymes could be used as targets for antibiotics and that the
present structures could act as useful guides for the design of specific inhibitors of S46 peptidases from
pathogenic organisms.
Figure 1: Subunit structure of DAP BII.
References
Garcia-Ruiz JM & Morena A, Acta Crystallogr. D Biol. Crystallogr. 50, 484-490 (1994)
Ogasawara W, et al., J. Bacteriol. 178, 6288-6295 (1996)
Sakamoto Y, et al., Sci. Rep. 4, article number 4977 (2014)
Suzuki Y, et al., Sci. Rep. 4, article number 4292 (2014)
Takahashi S, et al., J. Synchrotron Radiat. 20, 968-973 (2013)
115
Poster P-41
Abstract
Antimicrobial peptides bind to Escherichia coli ribosome
Weinert S1, Krizsan A1, Hoffmann R1, Sträter N1
1Institute
of Bioanalytical Chemistry, University of Leipzig, Leipzig, Germany
Antimicrobial peptides (AMPs) act as a part of natural immune systems against bacterial
infections (1). Examples for such peptides are apidaecin from Apis mellifera (honeybee)
(2) and oncocin from Oncopeltus fasciatus (milkweed bug) (3, 4). Both peptides belong to
the class of proline-rich antimicrobial peptides (PrAMPs) (5). The effect of AMPs against
bacteria should be used in the therapy of humans with antibiotic resistant bacteria
infections (1). For the treatment of humans with apidaecin and oncocin the peptides have
to be optimized concerning effectivity and toxicity (5).
For the optimization to medical use it is important to know the mechanism of action.
Therefore it will be a great help to determine the binding structure of an AMP and its
interaction partner. Interaction partners are proteins of bacteria, which are inhibited in
their function.
First results showed the 70S ribosome of Escherichia coli (E. coli) as an interaction partner
of peptides. The AMPs lead to a decrease of mRNA translation. oncocin has a greater effect
on the inhibition of ribosome function than apidaecin. Also both PrAMPs showed a very
good binding to the 70S ribosome of E. coli.
The dissociation constant of apidaecin 137 (Api137: gu-ONNRPVYIPRPRPPHPRL-OH, gu
is N,N,N',N'-tetramethylguanidino, O is L-ornithine) and oncocin 72 (Onc72:
VDKPPYLPRPRPPROIYNO-NH2, O is L-ornithine) is around 0,5µM for both. The peptides
show different regions and amino acids, which are important for their activity. Also the
peptide length has an influence of the binding strength to the ribosome.
References
(1) Boman HG, Annu. Rev. Immunol. 13, 61-92 (1995)
(2) Casteels P, et al., EMBO J. 8, 2387-2391 (1989)
(3) Schneider M & Dorn A, J. Invertebr. Pathol. 78, 135-140 (2001)
(4) Knappe D, et al., J. Med. Chem. 53, 5240-5247 (2010)
(5) Vaara M, Curr. Opin. Pharmacol. 9, 571-576 (2009)
116
Poster P-42
Abstract
pHrediction of crystallisation solutions
Kirkwood JS1, Hargreaves D2, O'Keefe S1, Wilson J1
1 York Centre for Complex Systems Analysis, University of York, Heslington, York, YO10
5GE, UK; 2 AstraZeneca, Discovery Sciences, Structure & Biophysics, Mereside, 50F49,
Alderley Park, Cheshire, SK10 4TF, UK.
In the crystallisation of biomolecules the pH of the experiment is one of the most
important parameters (McPherson, 1989, Newman et al., 2012). Prior knowledge of the
optimum pH to initialise crystallisation trials could drastically reduce the number of
experiments to be performed. For example, Kantardjieff and Rupp (2004) reported a
connection between the isoelectric point of a protein and the pH of the condition in which
crystallisation occurs. Although the pH of the solution can be measured accurately before
an experiment using a pH meter or determined using an acid-base indicator with a
spectrophotometer, this is unlikely to be the case for data deposited in the Protein Data
Bank (PDB) and used in data mining. Typically the pH is recorded as that of the buffer
component of the reservoir solution, which can be up to four pH units away from the
actual pH (Bukrinsky & Poulsen, 2001, Kirkwood et al., in press). If the pH is not recorded
accurately, it potentially becomes a misleading parameter in crystallisation data analysis.
In order to provide a more accurate post hoc estimate for the pH of crystallisation
experiments than the buffer pH, a neural network was employed to predict pH based on
the recorded concentrations of the different chemicals species in the solution. The pH for
5161 sets of conditions was determined using spectrophotometry and approximately twothirds of the data used to train the neural network. The remaining data was used as an
independent test set for which a correlation of 0.92 was achieved between the predicted
pH and spectrophotometric pH.
References
Bukrinsky JT & Poulsen JCN, Journal of applied crystallography 34, 533-534 (2001)
Kantardjieff KA & Rupp B, Bioinformatics 20, 2162-2168 (2004)
Kirkwood JS et al., Acta Crystallographica Section D: Biological Crystallography (in press)
McPherson A, Preparation and analysis of protein crystals (1989)
Newman J, Sayle RA & Fazio VJ, Acta Crystallographica Section D: Biological
Crystallography 68, 1003-1009 (2012)
117
Poster P-43
Abstract
Tackling crystallization issues of challenging targets: Three case
studies from Novartis Basel
Maschlej M, Honnappa S, Scherer-Becker D, Gossert A, Gutmann S, Hinniger A,
Izaac A, Koch E, Scheufler C
Novartis Institutes for BioMedical Research, Basel; CPC/ Structural Biophysics
At the interface of biology and chemistry, the Structural Biophysics Department at NIBR
Basel delivers structural information on target proteins and their interactions with ligands
to guide and support integrated lead finding and optimization.
Our success relies on the availability and quality of crystals. Therefore we focus on the
development of techniques to systematically screen and optimize protein crystals in an
automated high throughput manner. For challenging targets we often have to face specific
issues, which can occur at any step of the process from protein expression, purification to
crystal diffraction. For those difficult cases, we use the available knowledge to identify the
most suitable approach to obtain well-diffracting protein crystals.
The approaches can be addressed at different steps in the process:
• Cloning: limited proteolysis, entropy reduction, surface mutations
• Expression: solubility tags, co-expression with ligands in various hosts
• Purification: optimization and refolding, methylation, posttranslational modification
• Crystallization: screening methods, (matrix-) seeding, crosslinking, LCP
• Data collection: cryo optimization, in-situ diffraction tests
Here we present 3 techniques that allowed us to successfully determine the threedimensional structure of challenging protein targets.
Derewenda ZS, Acta Cryst. D67, 243-248 (2011)
Ronning D, Biochemistry 51, 9763−9772 (2012)
118
Poster P-44
Abstract
Automated protein crystal optimisation with TTP Labtech’s
dragonfly
Thaw P, Jenkins J, Cochrane G
TTP Labtech, Royston, UK
The ability to crystallise proteins, nucleic acids or macromolecular complexes pose
significant challenges to the protein crystallography community, from large scale
screening assays for the determination of initial crystallization conditions, screen
optimisation and final screen set-up.
Protein crystal optimisation is vital to ensure high quality X ray diffraction data for the
solving of high resolution structure. This process involves the set-up of a series of complex
screening combinations where the ratios of the individual components identified from
primary crystallisation studies are varied.
In order to reduce the effort and tedium of this process, TTP Labtech have introduced
dragonfly as an addition to their successful mosquito liquid handling portfolio for
crystallisation screening.
This poster demonstrates that “dragonfly” is a valuable, compact, low cost addition to the
crystallographer’s bench. It eliminates lengthy and complicated plate set-up at the
optimisation stage of crystallisation.
119
Poster P-45
Abstract
Random microseed matrix screening: A number of success stories
Moroz OV1, Blagova EB1, Friis EP2, Davies GJ1 and Wilson KS1
1Department
of Chemistry, York Structural Biology Laboratory, University of York, York,
UK;
2Novozymes A/S, Bagsvaerd, Denmark;
Random microseed matrix screening (rMMS) is a relatively new technique in
macromolecular crystallisation, first introduced by Ireton & Stoddard in 2004 (1) and
automated by D'Arcy et al (2). The crystals from a single hit, often of poor quality, are
crushed and the resulting seeding stock is introduced into the conditions of a new screen.
The concept is to separate nucleation from crystal growth, allowing new hits and better
diffracting crystals. In addition to self-seeding (with crushed crystals of the same protein),
cross seeding with homologous proteins is possible, as well as seeding of a complex with
crystals of one of the components (3,4).
We present here several examples of the successful use of rMMS, which confirm the
advantages of this powerful tool. The projects include both self-seeding and cross-seeding
for a number of proteases and their mutants, as well as a similar approach for amylases
and lipases set up by Mosquito (TTP Labtech) and Oryx (Douglas Instruments) robots.
References
1. Ireton GC & Stoddard BL, Acta Cryst. D60, 601-5 (2004)
2. D'Arcy A, Villard F, & Marsh M, Acta Cryst. D63, 550-4 (2007)
3. Obmolova G et al., Acta Cryst. D66, 927-33 (2010)
4. Shaw Stewart PD et al., Cryst. Growth Des. 11, 3432–3441 (2011)
120
Poster P-46
Abstract
Crystallization of green fluorescent protein for high-resolution Xray crystallography
Takaba K1, Takeda K1, and Miki K1
1Graduate School of Science, Kyoto University, Kyoto, Japan
High-resolution X-ray crystallography can be used to determine the detailed structures of
protein molecules, including the positions of hydrogen atoms and the distribution of
outer-shell electrons. However, radiation damage may result in serious problems that
prevent us from obtaining precise high-resolution structures of proteins. For collecting
the diffraction data with high quality, exposing multiple positions on a crystal could
reduce the radiation damage. A large crystal, which enables us to allocate multiple
irradiated positions, is required for high-resolution X-ray crystallography.
Green fluorescent protein (GFP) is used as a fluorescent reporter for various researches in
recent biology. In the chromophore of GFP, proton coupled electron transfer (PCET)
reactions occur with the fluorescence. Spectroscopic and computational studies on the
proton transfer in GFP have been reported. However, there have been no structural
studies where hydrogen atoms of the chromophore are determined.
By repeating micro- and macro-seeding methods, and controlling the temperature during
the crystal growth, we obtained an adequate size of crystals for high-resolution X-ray
crystallography (≥1 mm) of GFP variants. The S65T and T203I variants were used, which
have only the anionic and neutral chromophore, respectively. The mechanism of the
fluorescent reaction of GFP would be understood by comparing the hydrogen bonding of
these variants’ chromophore. The best X-ray diffraction data set of 0.94 Å resolution has
been collected so far. We are now optimizing the various conditions of crystal growth for
the large crystals needed for further high-resolution X-ray crystallography. We will
present the repeat-seeding and controlling the temperature for crystallization.
Suppressing the dose (<0.3 MGy) and cryo-cooling (at 25 K) with helium gas are
indispensable for preventing the destruction of chromophore by X-ray radiation.
Figure 1. A GFP crystal obtained with dimensions
of 2.1 x 0.3 x 0.3 mm3.
121
Poster P-47
Abstract
Haloalkane dehalogenases as subjects for structure-functional
studies
Kuta Smatanova I
Faculty of Science, University of South Bohemia in Ceske Budejovice, Ceske Budejovice,
Czech Republic andInstitute of Nanobiology and Structural Biology, Academy of Sciences
of the Czech Republic, Nove Hrady, Czech Republic
The protein crystallography is one of the methods used to study the protein structures
and to describe their function and mechanism. The most important and necessary
condition to used protein crystallography is obtaining of well-diffractable monocrystals.
Different crystallization techniques such as standard, advanced and alternative methods
are used to crystallize proteins and protein complexes. Nowadays more than 20 proteins,
protein complexes and their mutant variants from haloalkane dehalogenases family are
systematically studied. The main target is focused on research of proteins such as e.g.
DhaA from Rhodococcus rhodochrous NCIMB 13064, DbeA of Bradyrhizobium elkani
USDA94, LinB of Sphingobium japonicum UT26 or new haloalkane dehalogenases DpcA
from Psychrobacter cryohalolentis K5 and DmxA from Marynobacter sp ELB 17, etc. Finally
developed methods and obtained crystallization and crystallographic data are compiled
and results are published in scientific journals [1].
This research is supported by the GACR (P207/12/0775).
References
1. Chaloupkova R, et al., Acta Crystallogr. D Biol. Crystallogr. 70, 1884-1897 (2014)
122
Poster P-48
Abstract
Formamidase RCLECs: crystallographic studies and preliminary
use in acetohydroxamic acid production
Conejero-Muriel M1, Martínez-Rodríguez S2, Rodriguez Ruiz I3, Gavira JA1
de Estudios Cristalográficos, Instituto Andaluz Ciencias de la Tierra (IACT-CSIC-UGR), Armilla,
Granada, Spain. 2Departamento de Química-Física, Universidad de Granada, Granada, Spain.
3Institut de Microelectrónica de Barcelona (IMB-CNM, CSIC), Campus UAB, 08193 Bellaterra, Spain.
1Laboratorio
Formamidase (E.C. 3.5.1.49) catalyses the conversion of formamide into formic acid and ammonia with high
efficiency. The formamidase from Bacillus cereus also hydrolyses acetamide into acetic acid and ammonia,
and it is able to produce acetohydroxamic acid starting from both acetamide and acetic acid in the presence
of hydroxylamine (Soriano-Maldonado et al., 2011). Hydroxamic acids, in general, are weak organic acids
and find applications in biology and medicine. Particularly, acetohydroxamic acid is a potent and
irreversible inhibitor of bacterial and plant urease. Commercialised as Lithostat®, it is indicated as an
adjunct to antimicrobial therapy in patients with chronic urea-splitting urinary infection.
In this work, we have cloned, over-expressed and purified the formamidase from Bacillus cereus ATCC
14579. Crystallization screenings were performed in capillaries by counter diffusion technique. Crystal
improvement was achieved using different types of agarose gels and protein:precipitant ratios. Crystals
grown in different conditions were diffracted using X-ray synchrotron radiation and the structural 3D
models determined from crystals grown at pH ranging from 4.0 to 9.0 by Molecular Replacement. Crosslinked enzyme crystals (CLECs; Roy & Abraham, 2004; Arnoud et al., 2003) and Reinforced CLECs (RCLECs)
of formamidase have been produced both in capillaries and in microfluidic devices (microchips) (Vollrath C.
and Dittrich P.S., 2010) or in gels, respectively. While CLECs produced in microchips will reduce
dramatically the reactants volumes, RCLECs may allow an easy manipulation, preservation and
enhancement of the effectiveness of this solid catalyst. Preliminary results of the use and re-use of RCLECs
and the detection of activity on CLECs crystal produced in microchips devices will be shown. Our final goal
is to verify whether formamidase CLECs/RCLECs can be used for acetohydroxamic acid production at bench
scale.
References
1.
2.
3.
4.
Soriano-Maldonado P., et al., Applied and Environmental Microbiology 77, 5761–5769 (2011)
Roy J.J. & Abrahams T.E., Chemical Reviews, 104:9, 3705-21(2004)
Arnoud H. M., et al., The Journal of Biological Chemistry 278, 9052-9057 (2003)
Vollrath C. & Dittrich P.S, Microfluidics: Basic Concepts and Microchip Fabrication. In Bontoux N.,
Potier M-C., editors, Unravelling Single Cell Genomics : Micro and Nanotools, RSC Nanoscience &
Nanotechnology, 111–149 (2010)
123
Poster P-49
Abstract
Protein crystals architecture and diffraction quality controlled by special
additives
Hašek J1,2, Skálová T1, Dušková J1, Koval T2, Kolenko P2, Fejfarová K2, Stránský J1, Dohnálek J1,2
of Biotechnology, and 2Institute of Macromolecular Chemistry, ASCR, Praha
1Institute
Crystal architecture. Protein crystals represent a special state of matter. Majority of biomolecular surface remains
hydrated even in the crystalline state because protein crystals contain 30-80 % water retaining mostly its liquid state
properties. The biomolecules are held in their exact positions in crystal only by several repeating intermolecular
interfaces. The space arrangement and quality of these interfaces are critical for stability of the molecular scaffold and of
course for diffraction quality of the crystal.
Protein-protein adducts (PPA) formed in solution temporally (with short time-of-life) have important role in formation
of the condensed phase. As the adhesion interfaces remain in equilibrium with solution at any stage of crystallization
process (and even in the protein crystal), they depend crucially on any changes of solvent parameters (local
concentration changes of additives and other solvent components, dehydration, pH, etc.). The solvent variations affect
the crystallization negatively as a rule. However, when correctly used, they are an efficient tool to get uniform molecular
stacking leading to high diffraction quality. Protein molecules with a large molecular surface possess usually a number
of protein-protein adhesive patches. Thus, a number of various temporary protein-protein adducts (PPA) is usually
formed in solution. The adhesion propensity in different PPA can be largely controlled by special additives “Protein
Surface Active Molecules” (PSAM) [1,2,3]. Here, we summarize typical behavior of some PSAM (low affinity protein
surface binders). PSAM can act as adhesion competitors but also reversely as intermolecular adhesives (the ligand binds
to two or more neighbor molecules). The detailed knowledge of the PSAM action helps us to suppress the unwanted
adhesion modes. Improper choice of PSAM induces stresses, irregularities or a full loss of crystallinity. Used in
controlled manner, they offer an efficient tool for a design of the architecture of adhesion modes in the crystal,
optimization of adhesion patches and optimal balance of intermolecular forces ensuring high diffraction quality of the
crystal.
Protein crystallization. When all the incompatible adhesion modes are eliminated, the dominating adhesion mode
(DAM) ensures the desired dominance of a single PPA in sample. A complete elimination of all incompatible adhesion
modes leads to uniform stacking of molecules in the growing crystal. Thus, the scan over different crystallization
screens is in principle a search for conditions ensuring a dominance of a single DAM, which is necessary to get single
crystals for high quality structure determination. The concept of “protein-PSAM adducts” crumbled during
crystallization process is more complicated for imagination and requires detailed knowledge of rules governing a
unique formation of PPA. However, it provides a desired control over the crystal quality, explains many otherwise
enigmatic crystallization phenomena, and allows us determination of protein structures in different protein-protein
adhesion modes (different polymorphs).
Study of protein-protein interfaces in crystals is important to explain correctly the function of biomolecules. Protein
crystallizations with different DAMs gives a review of most important interaction modes and provide a realistic 3-D
view of weak protein-protein interfaces. It shows an important role of water and conformational changes of the
protein at different protein-protein interfaces. This information is hardly obtainable by other methods, namely if one
studies proteins in diluted solutions.
Proteins in their natural environment in living tissues act in concentrations exceeding many times their solubility
limits (water content often in range 50-80 %). It leads to limited diffusion and thixotropy properties. These
concentrations are well comparable with the concentration of biomolecules in crystals (water content in protein crystals
are in range 30-80 %). Thus, protein crystals are an excellent model for behavior of proteins in the living body
where they are in permanent contacts with other biomolecules, too. The molecularly overcrowded environment
leads in both cases to similar stresses leading to special side chain conformations and similar mobility restraints.
Inspection of the same protein structure in different crystalline environments (different crystal conditions [pH,
additives, solvent contents], different space groups, co-crystals, crystalline complexes) provide us more realistic insight
into the variability of protein structure and its behavior in living tissues.
The concept of protein-protein adducts and study of variety of protein structures from the Protein Databank lead to
better understanding of biological processes and can help in other biophysical methods.
The project is supported by BIOCEV CZ.1.05/1.1.00/02.0109 ERDF, and MSMT projects EE2.3.30.0029, LG14009, CSF
P302/11/0855.
References: [1] Hašek J, Z. Kristallogr. 23, 613-619 (2006); [2] Hašek J, Synchr. Radiation 18, 50-52 (2011); [3] Hašek J,
et al, Z. Kristallogr. 28, 475-480 (2011)
124
Poster P-50
Abstract
Calcium induced loss of rigidity in Myosin VIIa IQ5-post-IQ
continuous helix
Li J1, Deng Y1, Lu Q1, Zhang M1
1Division
of Life Science, State Key Laboratory of Molecular Neuroscience, Hong Kong
University of Science and Technology, Hong Kong SAR, China
Myosin VIIa is an unconventional myosin which plays important role in auditory and
visual systems. The structure and function of its highly charged helical region after IQ
motifs (post-IQ) have not been fully studied. We demonstrated that Myosin VIIa post-IQ
region forms a stable single helix (SAH) in both crystal and solution. This SAH together
with its preceding IQ5 constitutes a single rigid continuous helix with apo Calmodulin
(CaM) binding. We discovered that Myosin VIIa IQ5 interacts with Ca2+-CaM in a mode
distinct from IQ5-apo-CaM interaction. This Ca2+ induced conformational change breaks
the continuity and rigidity of IQ5-post-IQ continuous helix, serving as a potential
regulation mechanism for myosin force transduction. Integration of IQ5 and the SAH into
one entity extends the conventional CaM-modulated lever arm to the entire continuous
helix. We propose that this continuous helical structure and Ca2+ regulation property can
apply to other unconventional myosins including Myosin X and Myosin VI.
Modelled structure of the full length MyoVIIa: (A) Modelled structure
of MyoVIIa shows the contribution of SAH on lever-arm length. (B)
Working model of MyoVIIa as a tension sensor. (C) Working model of
MyoVIIa as a cargo transporter.
125
Poster P-51
Abstract
Expression of functional GPCRs for biophysical studies
Kubicek J1, Fabis R1, Dreyer M2, Langer T2, Hessler G2, Maertens B1
1
Cube Biotech GmbH, Alfred-Nobel-Str.10,40789 Monheim, Germany
Sanofi-Aventis Deutschland GmbH, Industriepark Hoechst, 65926 Frankfurt a.M.,
Germany
2
Human membrane proteins, in particular G-protein-coupled receptors (GPCRs), are
important drug targets. Precise characterization of ligands requires sufficient amounts of
pure and active protein that can be used in various assays to identify novel as well as more
specific drugs.
In this project, pharmaceutically relevant GPCRs were expressed, purified and tested for
activity. Notably, all proteins were expressed in their non truncated, wild-type form, with
different affinity tags added to the C- and N-termini. Bacterial and insect expression
systems were employed.
In a side-by-side analysis of different tag-based affinity chromatography methods, the
Rho1D4 tag showed highest specificity and yields e.g. compared to the polyhistidine tag.
In addition, an alternative affinity chromatography method was tested for its ability to
enrich active protein based on interaction with a specific ligand. Since free ligand has a
higher affinity compared to the matrix-bound ligand, it was possible to elute the protein
with the same ligand.
Biophysical measurements comparing ligand-binding properties of GPCRs purified from
different expression hosts and by different affinity matrices will be presented, and the
advantages and disadvantages of various methods will be discussed.
126
Poster P-52
Abstract
Crystallization of modified reaction centres of Rba. sphaeroides
Gabdulkhakov AG1, Fufina TY2, Vasilieva LG2 and Shuvalov VA2
1Institute
of Protein Research, Pushchino, Moscow region, Russia;
Pushchino, Moscow region, Russia
2Institute
of Basic Biological Problems,
Photosynthesis is the process of light energy transformation into a biologically usable form of chemical
energy. Light is first absorbed by light-harvesting antenna complexes and the energy is then passed to the
photosynthetic reaction centers (RC). RC of purple bacterium Rhodobacter sphaeroides is an integral
membrane pigment-protein complex which consists of three protein subunits and ten cofactors of electron
transfer. Cofactors are represented by four bacteriochlorophylls (BChl), two bacteriopheophytines, two
ubiquinones, a molecule of carotenoid and a non-heme iron atom. BChl structure includes a central Mg
atom, which considerably affects spectral properties of tetrapyrroles. The BChls microenvironment can
greatly affects their photophysical and redox properties and can be altered by site-directed mutagenesis.
Comparison of the crystal structures of mutant and wild type RCs can provide new information on
interactions between protein and cofactors of electron transfer.
Usually in bacterial RCs the fifth ligand of BChl Mg atom is histidine which is placed athwart to the plane of
the macrocycle. We study stability and properties of the dimer BChl structure using mutants with dislocated
position of the fifth Mg atom ligand. It is expected that this dislocation may change the distances between
BChls in the dimer P and thus to affect inter dimer interactions. Now we have structure of wild type RC at
2.3 Å resolution and several mutant structures from 2.4 Å to 3.0 Å resolutions [Vasilieva LG, et al., 2012;
Gabdulkhakov AG, et al., 2013]. Recently we obtain mutant form I(M206)H+H(M202)L RC. Spectral
properties and pigment composition of this double mutant RC was similar to I(L177)H+H(L173)L RC:
distinct QY absorption band of the dimer bacteriochlorophyll P disappears, and QY BChl band near 800 nm
broadens and shifts to the red. The pigment content of the mutant RC remains unaltered indicating secure
PA Mg atom coordination. [Vasilieva LG, et al., 2012]. Our attempts to crystallize RC I(M206)H+H(M202)L
encountered with difficulty of growing suitably large crystals of this unstable mutant complex. We got a
large number of small crystals, which in our opinion are suitable for use in X-ray pulse from free-electron
laser or serial crystallography synchrotron radiation [Stellato F,et al., 2014].
The authors would like to acknowledge financial support of RAS within the program “Molecular and Cell
Biology”, Russian Foundation for Basic Research (13-04-01148a).
Crystals of mutant form I(M206)H+H(M202)L RCs of the purple bacterium Rba. sphaeroides strain RV
References:
Vasilieva LG, et al., BBA 1817, 1407-1417, (2012)
Gabdulkhakov AG, et al., Acta Cryst. F69, 506-509, (2013)
Stellato F,et al., IUCrJ 1, 204-212, (2014)
127
Poster P-53
Abstract
Towards structural determination of the zinc finger domain of an
inner nuclear membrane protein
Vijayaraghavan B1, Hallberg E1
1Department
of Neurochemistry, Stockholm University, Stockholm, Sweden.
Samp1 is a transmembrane protein of the inner nuclear membrane, which is functionally
associated with centrosomes (Buch et al., 2009) and LINC complexes (Gudise et al., 2011).
During mitosis, Samp1 co-distributes with microtubules of the mitotic spindle (Buch et al.,
2009) suggesting a mitotic function.
To understand Samp1 functions in detail, a 3D structure of zinc finger domain of Samp1
would be extremely helpful. To determine Samp1 structure, we turned to thermophilic
fungus: Chaetomium thermophilum (Ct), which has ideal properties for structural studies,
because they tend to form stable 3D structure more easily (Amlacher et al., 2011). The
corresponding zinc finger domain of Samp1 was found in Ct. RNA was extracted from
frozen Ct cells followed by cDNA synthesis using reverse transcriptase. Different
constructs containing a zinc finger domain of Ct Samp1 (1-137, 1-180 and 1-180 - SGSGS320- 618) were amplified and cloned to histidine tagged pET-33b(+) vector. The
constructs Ct Samp1(1-137 and 1-180) were expressed in E. coli cells and found to be in
soluble form. Further, Ct Samp1(1-180) protein was purified using affinity
chromatography followed by gel filtration chromatography to homogeneity. The
purification of the remaining constructs will be performed and the purified protein
sample will be crystallised and structure will be determined by X-ray diffraction.
References
Jafferali MH et al., Biochim Biophys Acta, accepted
http://dx.doi.org/10.1016/j.bbamem.2014.06.008 (2014)
Gudise S et al., Journal of Cell Science 124, 2077-2085 (2011)
Buch C et al., Journal of Cell Science 122, 2100-2107 (2009)
128
Poster P-54
Abstract
Optimisation of crystal growth for structural biology
Budayova-Spano M1, Oksanen E1,2, Junius N1, Berzin C1, Vernede X1, Ferrer JL1
1Institute
de Biology Structurale, CEA-CNRS-UJF UMR 5075, Grenoble, France; 2current address: European
Spallation Source, Lund, Sweden
In X-ray protein crystallography it is relatively easy to obtain crystals of a protein, but it is considerably more laborious
to obtain crystals of sufficient size and quality of diffraction. Recording data from very small crystals requires
significantly longer measurement times and dozens (even hundreds) of crystals often must be used to collect a complete
set of data due radiation damage. The quality of data obtained is often lower than it could be with larger crystals.
In neutron macromolecular crystallography where the available neutron sources are weak, the crystal volume required
for a neutron data set is most often the factor limiting the more widespread use of this technique. If normal
hydrogenated proteins are used, a minimum crystal size of at least 1 mm3 is necessary in order to achieve sufficient
signal-to-noise ratios with the latest neutron sources (Blakeley, 2009). If perdeuterated proteins are used the minimum
crystal size can be as “little” as approximately 0.15 mm3 (Blakeley, 2009), but this is still (assuming an isometric crystal)
0.53 x 0.53 x 0.53 mm, i.e. of a size that most X-ray crystallographers no longer try to produce. However, even with these
large crystals, a single neutron diffraction dataset can take several days or weeks to collect. These and other technical
difficulties partly explain why only 70 neutron crystal structures (0.07 % of the total number) have been deposited in
the Protein Data Bank (http://www.rcsb.org/pdb) to date. Thus if neutron crystallography is to become a more routine
technique for the structural biology community then the ability to control crystal size must become more accessible and
routine for scientists working in the field. This is particularly important in order to push the limits of neutron
crystallography towards more challenging targets such as membrane proteins.
While the theoretical background of the crystallisation process is well established and studied with model systems
(Vekilov, 2005; Vekilov 2010), the principles are often difficult to implement in practice, as the level of supersaturation
cannot be controlled effectively in crystallisation setups using small enough amounts of precious protein. Novel devices
(Boudjemline et al., 2011; Meyer et al., 2012) attempt to address this issue with somewhat different strategies.
So, we developed a new crystallisation instrument that combines precise temperature control with real-time
observation through a microscope-mounted video camera (Budayova-Spano et al., 2007). In its latest development, this
crystal growth bench in addition to accurate temperature control also allows composition of the crystallisation solution
(e.g. precipitant concentration, pH, additive) to be controlled and changed in an automated manner (Budayova-Spano,
2011). Our approaches allow the rational optimisation of crystal growth (Oksanen et al., 2009) based on a
multidimensional phase diagram.
Typical crystal habits of a urate oxidase complex, here a
large crystal of urate oxidase complex with cyanide
(approximate dimensions 1.2 x 1.2 x 1 mm).
References
Blakeley M, Crystallography Reviews 15, 157-218 (2009)
Vekilov PG, Methods in Molecular Biology 300, 15-52 (2005)
Vekilov PG, Crystal Growth and Design 10, 5007-5019 (2010)
Boudjemline A, et al., Anal. Chem. 83, 7881-7887 (2011)
Meyer A, et al., Acta Crystallogr. F68, 994-998 (2012)
Budayova-Spano M, et al., Acta Crystallogr. D63, 339-347 (2007)
Oksanen E, et al., J. R. Soc. Interface 6, S599-S610 (2009)
Budayova-Spano M, Patent EP 117730945 (US13821053, JP2013528746, AU2011303702), UJF (2011)
129
Poster P-55
Abstract
Crystallisation of ferredoxin/flavodoxin-NADP(H) oxidoreductases,
flavodoxins, ferredoxins and redox partners in Bacillus cereus
Lofstad M1, Gudim I1, Skråmo S1, Hammerstad, M1, Røhr ÅK1, Tomter AB1,
Andersson KK1, Hersleth H-P1
1Depatment
of Biosciences, Section for Biochemistry and Molecular Biology, University of
Oslo, Oslo, Norway
Ferredoxin/flavodoxin-NADP(H) oxidoreductases (FNRs) catalyses the reversible redox
reaction between ferredoxins (Fds) or flavodoxins (Flds) and NAD(P)+/NAD(P)H. In
oxygenic photosynthetic organisms FNRs catalyse the reduction of NADP+ by
photosynthetically reduced Fd, while in many heterotrophs, FNRs catalyse the NADPHdependent reduction of Fd/Fld to provide subsequently reducing power to different
Fd/Fld-dependent enzyme systems. In Bacillus cereus there are three annotated FNRs that
can be redox partners for three Flds and two Fds. We have without tags purified and
crystallised several of these redox proteins in Bacillus cereus, and some of their redox
partners. A goal of the project is to understand the recognition, selectivity and flexibility of
these FNRs, Flds and Fds with respect to each other and their redox partners.
References
Skråmo S et al., Acta Cryst. F70, 777-780 (2014)
Hammerstad M et al., ACS Chem. Biol. 9, 526-537 (2014)
Røhr ÅK et al., Angew. Chem. Int. Ed. 49, 2324-2327 (2010)
130
Poster P-56
Abstract
Development and application of a DLS-based solution circulating
crystallization apparatus
Miyatake H1, Yano T2, Shiroma R2, Fukumoto N2, Matsumoto K3, Kobayashi M2,
Maezawa S2, Kuriyama N2
1RIKEN
Global Research Cluster Collaboration Promotion Unit, Saitama, Japan
Co., Ltd., Kyoto, Japan
3DFC Co., Ltd., Kyoto, Japan
2YMC
We already reported the prototype of new concept crystallization apparatus based on the
DLS measurement (Miyatake H et al.). In the apparatus, solution composition in the
solution circuit is under control based-on the DLS measurement. In the crystallization
procedure of the apparatus, protein and/or precipitant concentrations are gradually
increased keeping the dispersity of protein as monodisperse as possible. Such a way of
real-time controlling of protein aggregation in nano-level will prepare better crystallinity
and larger size of crystals. The former experimental crystallization apparatus grew highquality HEWL crystals which diffracted x-ray to 1 Å resolution. In addition, the apparatus
grew larger HEWL crystals up to ~2 mm, which will be useful to prepare larger crystals
suitable for neutron diffraction crystallography.
We have collaborated with a company, YMC Co., Ltd., to develop a sophisticated
crystallization apparatus for customer use level. In the apparatus (Figure 1), all the units
for measurement will be compactly placed together on the body table. During
crystallization, they are integrally controlled by a Windows PC. We will present the
current status of the new crystallization apparatus which is under developing.
DLS device
Figure 1: A conceptional drawing of a new solution
circulating DLS-based crystallization apparatus. All
of the measurement units; CCD camera,
conductivity meter, UV meter, pH meter, and DLS
device are aligned in the solution circuit. The
protein crystallization cell is also inserted into the
solution circuit, which is constantly monitored by
the CCD camera. Protein and other buffer solutions
are injected via the multi-valve positioner. The
solution in the circuit is compulsory circulated by
the peristaltic pump.
References
Miyatake H, et al., Crystal Growth & Design 12, 4466-4472 (2012)
131
Poster P-57
Abstract
Preparation of protein nano crystals and scoring by dynamic light
scattering on the basis of their radius distribution pattern
Schubert R1,2‡, Meyer A3‡, Künz M1, Dierks K3, Perbandt M1,2, Betzel C1,2
1Institute
of Biochemistry and Molecular Biology, University of Hamburg, Hamburg,
Germany; 2Hamburg Centre for Ultrafast Imaging, Hamburg, Germany; 3XtalConcepts
GmbH, Hamburg, Germany
‡ Authors contributed equally to this work
Latest techniques and advanced inhouse developed methods to precisely monitor crystal
growth and to optimize the preparation of crystalline particles, too small to be observed
by light microscopes, will be presented[1]. Growth and preparation of high quality microsized crystals optimal for data collection experiments at modern micro-beam insertiondevice synchrotron (SR) beamlines and growth of nano crystals required for data
collection at Free-Electron-Laser (FEL) beamlines is a new and challenging task. In
principle nano crystals can be formed quite readily by controlled driving of a protein
solution into supersaturation, however, techniques are required to control and score the
process online.
Therefore new and advanced methods to prepare and score 3D micro- and nano crystal
suspensions, most suitable for X-ray and electron diffraction experiments need to be
established. During crystallization biomolecules pass through different stages[2], which are
continuously characterized on the basis of dynamic laser light scattering (DLS). For a
successful crystallization, the knowledge and characterization of the concentration and
time dependent processes is of fundamental importance. In particular, the detection of the
first nucleation is essential for the further crystallization success and can be visualized by
electron microscopy, but not by conventional light microscopy methods[3,4]. We were able
to identify a DLS "fingerprint" which reports the successful growth of nano crystals with a
size of 50 nm to 300 nm. For the identification and scoring of this fingerprint different
samples were analyzed by transmission electron microscopy.
The data confirm that DLS is most suitable to distinguish between protein aggregates and
crystalline particles on the basis of the radius distribution of the particles in solution, even
if both have a similar size. This enables us to evaluate the formation of nano crystals in the
crystallization solution without time consuming electron microscopy investigations.
[1] Meyer A, et al., Acta Cryst. F68, 994-998 (2012)
[2] Vekilov PG, Nanoscale, 2, 2346-2357 (2010)
[3] Gomery K, et al., Microsc. Microanal. 19, 145–149 (2013)
[4] Stevenson HP, et al., PNAS, 111, 8470–8475 (2014)
132
Poster P-58
Abstract
Crystallization techniques and services at the Biocenter Finland
crystallization facility
Kogan K1, Mäki S1., Kestilä VP1, Goldman A2, Kajander T1
1Institute
2Division
of Biotechnology, University of Helsinki, Helsinki, Finland
of Biochemistry, Department of Biosciences, University of Helsinki, Helsinki,
Finland
Protein crystallization has become automated to variable extents during last tenfifteen years with the emergence of structural genomics and other high-throughput
demanding structural biology research efforts. While many structural biology groups and
centres have equipment for their own purposes, our goal as a national service unit, like in
many other much larger centres, is to provide users and customers with a wider range of
services and with a full package of options.
At the moment we can offer domestic, international and industrial users
crystallization services with several options. The basic core set includes standard sitting
drop setups with TTP labtech mosquito and imaging with Rigaku minstrel UV at room
temperature (+22°C) and Explora Nova colour imaging station at +10°C. We have
developed our own viewer, PixRay (Kestilä et al, unpublished) for remote inspection and
scoring of imaging results through a web browser. Other main crystallization options
include micro-seeding/re-screening with seeding and options to setup LCP-plates on the
Mosquito and have strong experience on membrane proteins (e.g. Kellosalo et al. 2012).
We also offer thermofluor screening and synchrotron data collection service at ESRF and
Diamond light sources. Current developments include more unique opportunities; in
particular in collaboration with the Institute for Molecular Medicine Finland (FIMM) we
can offer a service for acoustic dispensing of limited-size samples with drop-sizes down to
20-25 nl. We also have options for fluorescent trace-labeling for screening e.g. for protein
complex crystals. Finally, we are also developing protein engineering strategies to enhance
crystallizability of proteins e.g. through sequence conservation-based approaches; a caseexample will be presented.
133
Poster P-59
Abstract
Understanding the Ubiquitin activating enzyme1
Misra M, Schäfer A, Schindelin H
Rudolf Virchow Zentrum, University of Würzburg, Würzburg, Germany
Post-translational modification of proteins by ubiquitination is a widely utilized phenomenon to alter the protein
function and fate in eukaryotes. The ubiquitin pathway is highly conserved throughout eukaryotes and consists of
well-established enzymatic system classified as E1, E2 and E3s. E1 corresponds to ubiquitin activating enzyme which
has three major activities: adenylation of C-terminal of ubiquitin in ATP dependent manner, transfer of activated
ubiquitin to its catalytically active cysteine residue present at the second catalytic cysteine half (SCCH) domain by
thioesterification reaction and transfer of ubiquitin from this site to E2 enzyme. E2s (ubiquitin conjugating enzymes)
then take up ubiquitin from E1 again through their catalytic cysteine residue and then pass it on to the E3 enzymes
(ubiquitin ligating enzymes) which either act as scaffolding protein between target protein and E2 or take up ubiquitin
from E3 and then ubiquitinylate the substrate on their lysine residue facilitated by an isopeptide bond between εamino group of Lysine and C-terminal glycine of ubiquitin. There are two E1 enzymes, nearly thirty E2s and several
hundreds E3s reported in humans. These enzymes participate in regulating the function and localization of numerous
cellular proteins through ubiquitin, enhancing the complexity of cellular proteome.
ATP
Ub(a)
(A)
(B)
(C)
Fig: Uba1 (Sacchromyces cerevisiae) in its different functional states; (A) ubiquitin bound at active adenylation domain
(PDB entry: 3CMM); (B) ATP bound form of Uba1 (unpublished) (C) ternary complex with activated ubiquitin present
at adenylation domain and another ubiquitin attached to the catalytic site cysteine via thiolester bond (PDB entry:
4NNJ).
Our group is interested in characterising structure to function relationship of Ubiquitin activating enzyme 1 (Uba1).
The UBA1 gene of the yeast S. cerevisiae encodes a 114kd Ub-activating enzyme which is essential for cell viability.
Uba1 plays a crucial role in activating and transferring ubiquitin to different E2s. We have been able to capture
different functional states of Uba1 which provide us great deal of information about the structural mode of action of
this enzyme. The targeted regulation of proteins by the ubiquitin pathway regulates numerous cellular processes
including protein degradation, cell proliferation, signal transduction, apoptosis, DNA repair, transcriptional regulation,
antigen processing, receptor modulation as well as endocytosis. The ubiquitin system controls the turnover and
biological function of many proteins within cell and alterations in the process can contribute to cancer progression,
neurodegenerative disorders and pathogenicity associated with microbes. The significance of ubiquitination as posttranslational modification and its impact on various cellular processes makes it important to understand the
mechanism of transfer of ubiquitin and we show the very first step of the ubiquitin pathway with relevant structures
as proof of concept.
References:
Haas AL et al., J. Biol. Chem. 257, 2543-2548 (1982)
Haas AL, Warms JV and Rose IA, Biochemistry 22, 4388-4394 (1983)
Hershko A and Ciechanover A, Annu. Rev. Biochem. 67, 425-479 (1998)
Lee I and Schindelin H, Cell 134, 1-11 (2008)
Schäfer A, Kuhn M and Schindelin H, Acta Crystallogr D Biol Crystallogr. 70, 1311-20 (2014)
134
Poster P-60
Abstract
Crystallization of uridine phosphorylase from Yersinia
pseudotuberculosis and it’s X-ray structure at 1.4 Å resolution
Balaev VV1, Lashkov AA1, Gabdoulkhakov AG1, Mikhailov AM1, Betzel C2
1Institute
of Crystallography Russian Academy of Sciences, Moscow, Russian Federation; 2Institute of Biochemistry and
Molecular Biology, University of Hamburg, Hamburg, Germany
Antibiotics – inhibitors of dihydrofolate reductase are used for treatment of acute infectious deseases,
caused by bacteria of Yersinia sp., family Enterobacteriae. Uridine phosphorylases take part in metabolism of these
drugs and, in particular, cytostatic antibiotic trimethoprim in the cells of these bacteria. Uridine phosphorylase
structures are essential for the determination of structural origins of interaction between these enzymes and
trimethoprim. Crystals of uridine phosphorylase Yersinia pseudotuberculosis (YptUPh), applicable for X-ray analysis,
were grown. The dynamic light scattering spectrometer SpectroSizeTM 300 (X-tal Concepts) was used as it is useful for
detection of size inhomogeneities. Crystallization solution optimization was held by means of laser dynamic light
scattering on the experimental facility X-tal Controller in laboratory of structural biology of infections and
inflammations University of Hamburg (DESY, Hamburg, Germany). Crystallization was held by means of vapor diffusion
method at 12°C (reservoir solution (50 µl): 0.1 M Tris-HCl, 5 % w/v PGA-LM , 20 % w/v PEG 2K MME, pH 7.8; Drop
solution: 5μl YptUPh (10 mg/ml in10mM Tris–HCl (pH 6.5)) and 5μl of reservoir solution). In order to obtain larger
crystals the following methodology was used: crystallization drop was equilibrated for 12 hours against reservoir
solution, then a piece of glass with a drop was transferred to a reservoir containing 400 μl of H2O and then back to a
resevoir solution. Large crystals were formed in a droplet after 1-3 days. X-ray data sets from these crystalls were
obtained on the station P11 (PETRA III, EMBL/DESY, Hamburg, Germany), what allowed to solve YptUPh structure at
atomic resolution. Spatial structure of uridine phosphorylase from Yersinia pseudotuberculosis was solved, refined and
deposited to RCSB PDB at 1.40 Å resolution (ID PDB: 4OF4, R-work=15.2%, R-free=18.4%, r.m.s.d.(bond
length/angle)=0.007Å/1.170˚, DPI=0.062 Å, Ramachandran statistics: Most favored regions: 98.30%; Allowed regions:
1.32%).
Spatial structure of YptUPh complexed with trimethoprim (TOP) and its modification 53I, which has higher
activity than other TOP modifications, was determined by means of molecular docking. Negatively-charged TOP
trimethoxybenzyl group is located in the area of primarily positive amino acid residues of YptUPh phosphate-binding
site. As for modification 53I, pyrimidine diamine group, which is positively charged as well, is located in phosphate
binding site. Hence, the binding energy between the enzyme and 53I is lower than between the enzyme and TOP.
Pyrimidine diamine group of TOP and dimethoxybenzyl group of 53I are localized in ribose-binding site. Trimethoprim
binds with the enzyme by means of four hydrogen bonds: O3_TOP – 3.2 Å – OG1_Thr94/A, O3_TOP – 3.5 Å – N_Thr94/A,
N1_TOP – 3.4 Å – O_Thr94/A, N4_TOP – 3.4 Å – SD_Met197/A. 53I forms following hydrogen bonds: O2_53I – 3.6 Å –
OE1_Gln226/A, O3_53I – 3.1 Å – NH1_Arg168/A, N4_53I – 2.8 Å – O_Thr94/A.
Molecular dynamic simulation revealed stability of both complexes. N1-glycosidic bond, which is present in
substrates of uridine phosphorylase, is absent in TOP and 53I molecules. Consequently, the reaction of their cleavage
cannot pass, and only their buffering is possible. This leads to the reduction of the antibiotic (and its modification)
effective concentration in bacterium cell. Thus, the treatment of infectious deseases, caused by bacteria of Yersinia sp.,
becomes more complicated. In future, rational drug design is planned in order to obtain a drug, binding target enzyme,
and not buffering by uridine phosphorylase.
(a)
(b)
Binding of trimethoprim molecule (a) and its modification(b) with active site amino acid residues of Yersinia
pseudotuberculosis uridine phosphorylase as a result of molecular dynamics simulation.
135
Poster P-61
Abstract
Purification, characterization and crystallization of Scytalidium
thermophilum xylanase
Sutay Kocabas D1, Tur E1, Guder S2
1Department
of Food Engineering, Karamanoglu Mehmetbey University, Karaman, Turkey;
Chemistry, Karamanoglu Mehmetbey University, Karaman, Turkey
2Department of
Xylanases are hydrolytic enzymes responsible for xylan depolymerization. Xylan is the complex
polysaccharide of the plant cell wall mainly consisting of D-xylose as the monomeric unit and it is
the most abundant non-cellulosic renewable polysaccharide on the earth (Beg et al. 2001, Dhiman
et al. 2008). Fungal xylanases are favorable at industrial scale such as animal feed production,
manufacture of bread, food and drinks, pharmaceutical and chemical applications, textiles, pulp
and paper production (Polizeli et al. 2005). Microbial xylanases are preferred biocatalysts in
industry due to their high substrate specificity, mild reaction conditions, conservation of substrate
and insignificant side product formation (Kulkarni et al 1999).
S. thermophilum xylanases were produced as described previously with a slight modification
(Sutay Kocabas et al. 2008). Ground corn cob particles (<2 mm sizes) were used at 20 g/l
concentration in the main culture as the carbon source. By using two-step chromatography
technique including gel filtration and anion exchange, low molecular weight xylanase was purified
ca. 21.8 fold with 9.6% recovery to apparent homogeneity as demonstrated by SDS-PAGE. The
enzyme has a molecular weight of 21 kDa and an isoelectric point of pH 8.6. Optimum temperature
and pH values of purified xylanase were found to be 65°C and 6.5, respectively. Xylanolytic
activity was most stable at pH 7.0 and 40°C. Purified xylanase showed the maximum specificity
towards beechwood xylan and wheat bran among investigated commercial and lignocellulosic
substrates.
Xylanase crystal growth was screened using ready-to-use kits from Hampton Research (USA) by
sitting drop vapor diffusion method. After dye test of the observed crystals, crystal forming
conditions were optimized in 24-well plates by hanging drop vapor diffusion technique at 18°C
by mixing an equal volume of the protein with reservoir solution. So far, best crystals, having
potential for X-ray structure studies, were obtained using ammonium citrate dibasic (1.6-1.8 M)
and sodium acetate trihydrate (0.10-0.12 M) at 18°C at 3 mg/ml protein concentration. An
example of a xylanase crystal is shown in Figure 1.
(a)
(b)
Figure 1. Crystal of S. thermophilum xylanase, (a) original crystal (b) crystal after dye treatment
References
Beg QK et al., Appl. Microbiol. Biotechnol. 56, 326-338 (2001)
Dhiman SS et al., Bioresources 3, 1377-1402 (2008)
Kulkarni N et al., FEMS Microbiol Rev 23, 411-456 (1999)
Polizeli MLTM et al., Appl. Microbiol. Biotechnol. 67, 577-591 (2005)
Sutay Kocabas D et al., Appl. Microbiol. Biotechnol. 79, 407-415 (2008)
136
Poster P-62
Abstract
Crystallization of complexes of VchUPh using dynamic light
scattering method and specifity of uridine phosphorylase from
Vibrio cholerae
Lashkov AA1, Prokofev II1, Gabdoulkhakov AG1, Betzel C2, Mikhailov AM1
1Shubnikov
2Institute
Institute of Crystallography, Russian Academy of Sciences, Moscow, Russia;
of Biochemistry and Molecular Biology, University of Hamburg, Hamburg,
Germany
Nucleoside re-synthesis processes in the eukaryotic and prokaryotic cells depend on activity of
pyrimidine phosphorylases, possessing both broad and narrow specifity. Research of different
aspects of uridine phosphorylase specifity is essential for the optimization of enzymatic methods
of nucleoside derivative obtaining and for rational drug design. Spatial structures of uridine
phosphorylase from pathogenic bacterium V. cholerae (VchUPh) complexed with substrates of
direct and reverse reactions were defined in order to investigate substrate specifity structural
origins of this enzyme.
In order to check for protein solution homogeneity before crystallization the spectrometer of
dynamic light scattering SpectroSizeTM 300 (X-tal Concepts) was used. Crystallization solution
composition was optimized, using automatic single-drop optimization method controlled by laser
dynamic light scattering on the experimental facility X-tal Controller in laboratory of structural
biology of infections and inflammations University of Hamburg (DESY, Hamburg, Germany). As a
result, the high quality crystals were grown during short time in ground conditions. This allowed
to obtain high resolution X-ray data sets, which were collected on DORIS and PETRA III
synchrotrons (EMBL/DESY, Hamburg, Germany). By means of X-ray analysis there were solved
structures of complexes: VchUPh+thymidine (1.29 Å, Rwork= 17.6%, Rfree= 21.0%, ID PDB:
4LZW), VchUPh+thymine (1.25 Å, Rwork= 11.5%, Rfree= 14.7%, ID PDB: 4OGL), VchUPh+uracil
(1.91 Å, Rwork= 17.2%, Rfree= 21.6%, ID PDB: 4OEH).
It was shown that VchUPh substrate preference to uridine is influenced by ligand 2’-hydroxy
group binding amino acid residues of ribose-binding site of the enzyme. This leads to the fixation
of uridine ribose part in configuration more predisposed for the reaction in contrast with
thymidine and to the increase of enzyme to uridine affinity. Thymidine 5-methyl group does not
interact with enzyme amino acid residues and can affect reaction speed only by means of positive
inductive effect on pyrimidine ring of the direct reaction ligand.
Interaction of thymine 5-methyl group with Ile219, Ile220 leads to changes in this ligand position
in contrast with uracil by 0.5 Å. In case of nucleoside synthesis reverse reaction this circumstance
makes an obstacle for thymine molecule nucleophile attack on the second substrate (ribose-1phosphate and 2-deoxyribose-1-phosphate). Thus various effect of 5-methyl radical of pyrimidine
ring on direct and reverse reaction catalysed by VchUPh is shown.
The reported study was partially supported by budget financing of Institute of Crystallography
Russian Academy of Sciences and by RFBR, research project No. 14-04-00952 a.
137
Poster P-63
Abstract
Recrystallization: A method to improve quality of protein crystal
Hou H, Tao HR, Yin DC*
Key Lab for Space Bioscience & Biotechnology, School of Life Sciences, Northwestern Polytechnical
The three dimensional structures of protein give the basis for the function of proteins, which is the reflection
of the life process. Therefore determining the structure of protein molecules is crucial. Nowadays, more than
89% of the protein structures in Protein Data Bank (PDB) have been determined by X-ray diffraction.
However, protein crystals of high quality for X-ray diffracting are still a bottleneck for structure determine.
In order to conquer this problem, numerous methods have been studied to improve the crystal quality.
Among them, chemical methods, such as increasing crystal screening conditions and add various chemical
small molecules to crystal growth condition, have been studied in searching for more suitable crystal
growth condition to cover the potentially useful crystallization space. Additionally, the effect of physical
environment, like microgravity, magnetic field, temperature and electrical field on protein crystallization
have been researched for seeking appropriate techniques to improve protein crystal quality. Moreover,
other important approaches, such as protein engineering which uses recombinant or mutagenesis
techniques to modify the protein to enhance its crystallizability, and microseeding which has been critical
for obtaining diffraction-quality crystals for many structures, are widely used in structure biology.
All of the abovementioned methods have potentially positive effects on protein crystal quality. However,
they all need more chemical conditions, additional physical environment, or molecular cloning technology. It
is important to invent a “native” method to improve protein crystal quality. In this paper, recrystallization -a
traditional method which does not change crystallization condition or does not need any other techniques,
was conducted to compare crystal quality before and after every recrystallization step. Five different
proteins were crystallized and crystal quality was checked by X-ray diffraction analysis. In this report, we
show that recrystallization indeed improve the crystal quality, and that the crystals which grow after
recrystallization step exhibit better morphology before recrystallization, indicating that the recrystallization
can be adopted in many protein crystallization optimization strategies.
Figure 1. A comparison of the resolution limit of protein crystals grown under different environments. The
data were averaged and normalized based on the resolution of the crystals grown in the control.
Acknowledgements
This work was supported by the National Natural Science Foundation of China (Grant No. 31200551), the PhD programs
Foundation of the Ministry of Education of China (Grant No. 20126102120068), and the Natural Science Basic Research
Plan in Shaanxi Province of China (Grant No 2013JQ4032)
Reference
Zhang CY et al., Crystal Growth & Design,. 8(12), 4227-4232 (2008)
Taleb M et al., Journal of Crystal Growth,. 200(3-4), 575-582 (1999)
Sazaki G, Moreno A, and Nakajima K, Journal of Crystal Growth,. 262(1-4), 499-502 (2004)
Derewenda ZS, Methods, 34(3), 354-63 (2004)
Longenecker KL et al., Acta Crystallogr D Biol Crystallogr, 57(Pt 5), 679-688 (2001)
Ireton GC and Stoddard BL, Acta Crystallogr D Biol Crystallogr, 60(Pt 3), 601-605 (2004)
138
Poster P-64
Abstract
The open national macromolecular crystallization facility at the
MAX IV laboratory
Håkansson M1,3, Kimbung R1, Welin M1, Otten H2,3,4, Appio R3, Fredslund F3,
Thunnissen MM2,3, Logan DT1,2,3
1SARomics
Biostructures AB, Lund, Sweden; 2Department of Biochemistry and Structural Biology,
Lund University, Lund, Sweden; 3MAX IV Laboratory, Lund University, Lund, Sweden; 4Department
of Biology, Lund University, Lund, Sweden
The MAX IV Laboratory (Eriksson et al., 2011, Leemann et al., 2009) in Lund hosts a highly
efficient national facility for high-throughput, nanovolume characterization and crystallization of
biological macromolecules. The facility enables users to carry out a wide variety of robot-assisted
crystallization experiments. The success rate of initial crystallization trials is maximized by
sampling many hundreds of conditions using minimal sample volumes. The facility can also be
used for software assisted design and implementation of optimization screens to produce crystals
suitable for data collection.
The proximity of the crystallization lab to the macromolecular crystallization beamline I911
(Cassiopeia; Ursby et al., 2013) allows for rapid and flexible evaluation of obtained crystals,
including direct scanning of crystallization plates in the X-ray beam.
In addition, users can characterize proteins and other biomolecules to determine and maximize
the chance of obtaining crystals. Alongside analysis of particle size distribution using dynamic
light scattering (DLS), one can screen for conditions that maximally stabilize the proteins using the
thermal shift assay (also known as DSF or Thermofluor).
The crystallization facility is open to academic users from the whole of Sweden (other countries
upon request via http://www.maxlab.lu.se/crystal) for modest access fees. The facility
acknowledges financial support from the Knut and Alice Wallenberg Foundation (SWEGENE
programme), the Erik & Maja Lundqvist Foundation, the Carl Tesdorpf Foundation, the Swedish
Research Council and the Natural Science Faculty of Lund University.
The National Crystallization Facility at the MAX IV Laboratory is managed by SARomics
Biostructures on behalf of the MAX IV Laboratory.
References
Eriksson M et al., IPAC'11 (2nd International Particle Accelerator Conference), 3026-3028 (2011)
Leemann SC et al., Phys. Rev. ST Accel. Beams 12, 120701 (2009)
Ursby T et al., J. Synchrotron Radiat. 4, 648-653 (2013)
139
Poster P-65
Abstract
Promoting protein crystallization using a cross-diffusion
microbatch method
Cao HL1,2, Chen RQ1, Liu YM1, Yin DC1*
1Key
Lab for Space Bioscience & Biotechnology, School of Life Sciences, Northwestern Polytechnical
2Institute of Basic and Transitional Medicine, Xi’an Medical University, Xi’an, Shaanxi, PR China
Growing diffracting quality crystals has been a bottleneck to determine protein structure using X-ray
diffraction (XRD). It is the key step to find a lead to obtain protein crystals. In this presentation, a new
method called the cross-diffusion microbatch (CDM) method is developed, which aims to efficiently increase
the chance of obtaining protein crystals. A very simple crystallization plate was designed for the CDM
method, in which all crystallization droplets are in one sealed space. As a result, various volatile components
and water can diffuse from one droplet into any other droplet via vapour diffusion, as shown in Fig. 1(a).
In this presentation, we will show the crystallization screening of 15 proteins using CDM method and the
comparison with conventional sitting-drop vapour-diffusion (SDVD) method. Fig.1 (b) shows statistical
comparison of the average normalized screening hits between the CDM method and the SDVD method (n =
14).
(a)
(b)
Fig.1 (a) Schematic of the crystallization plate for the CDM method; (b) statistical comparison of the
average normalized screening hits between the CDM method and the SDVD method (n = 14). The number of
hits was normalized on the basis of the data for the CDM method.
It was found that the CDM method can significantly increase the number of crystallization screening
hits compared with the SDVD method. The results indicate that the CDM method can dramatically increase
the chances to obtain protein crystals at a lower cost as well as a simple and easy procedure and could be a
potentially powerful technique in practical protein crystallization screening.
Acknowledgements
This work was supported by the National Basic Research Program of China (973 Program; grant No.
2011CB710905), the National Natural Science Foundation of China (grant Nos. 31170816 and 51201137),
the Doctorate Foundation of Northwestern Polytechnical University (grant Nos. CX201120 and CX201327)
and the Ministry of Education Fund for Doctoral Students Newcomer Awards, China.
References
Chen RQ et al, Acta Cryst. D 70, 647-657 (2014)
Cao HL et al., Acta Cryst. D 69, 1901-1910 (2013)
Lu HM et al., Rev. Sci. Instrument 79, 093903 (2008)
Yin DC et al., J. Cryst. Growth 310, 1206-1212 (2008)
140
Poster P-66
Abstract
Optimization of trace fluorescent labeling derived cryptic screening
leads with ionic liquids
Pusey ML, Barcena J, Morris M, Yuan Q, Ng J
iXpressGenes, Inc., Huntsville, AL 35806
Protein crystallization is treated as a binary process; yes there are crystals, no there are
not. The apparent negative outcomes are immediately dismissed from further
consideration. We use trace fluorescent labeling (TFL) as a means of rapidly identifying
crystals during screening. The method involves the covalent labeling of between 0.1 to
0.2 % of the protein molecules with a fluorescent probe. All results obtained to date show
that labeling below 1 % does not affect the nucleation rate or diffraction data quality.
Identification of crystalline outcomes is based on intensity; the protein packing density is
highest in the crystalline form which fluoresces more brightly than other precipitated
forms. We are using a 6 plate, 3 TFL+ and 3 TFL-, experimental approach with a single
screen to test the effects of TFL on crystal nucleation. We find that there are many
outcomes where the fluorescent images have regions or spots of high intensity, but no
corresponding (semi)crystalline structures are apparent with standard transmission
microscopy. The governing paradigm for TFL is that intensity = structure. From this we
hypothesized that these are likely lead conditions and are testing that hypothesis with
optimization screening. To date the number of TFL+ hits in the initial screen data is
slightly higher than for the TFL- hits. However, these additional leads can only come from
the TFL+ screening plates. Our standard optimization method uses capillary counter
diffusion (CCD). Overall success rates for optimization of the TFL derived leads are
~40 % from CCD experiments. However, we are now exploring the use of ionic liquids
(IL’s) as crystallization optimization additives. Seven commercial off the shelf IL’s are
being tested in this first round of experiments, with the IL’s used at 0.1 M final
concentration. The proteins employed in this study are not the usual models, but rather
derived from several ongoing research projects in this laboratory. Based upon the results
to date the IL-based optimizations are at least equivalent to those using CCD, with the
added benefit that the IL optimizations can be set up more quickly. The results show IL
structure dependence for the outcome, suggesting that the IL structures can be modified
to further improve their effectiveness. However, with the same protein different IL’s
worked best with different screen conditions, suggesting a link between the precipitant
and IL. Independent of the approach employed, even proteins that did not give crystals in
the initial screen have given crystals after optimization of the TFL-derived leads,
indicating the potential utility of this approach.
141
Poster P-67
Abstract
Crystallization of C-terminal domain (CTD) of Myosin IB from
Entamoeba histolytica
Gautam G, Abdulrehman SA, Gourinath S
School Of Life Sciences, Jawaharlal Nehru University, New Delhi, India
Myosin IB, the only unconventional myosin motor present in E. histolytica had been found to be
localized with early and late phagosomes as well as internalized phagosomes in E. histolytica cells, activated
for erythrophagocytosis. Over expression of Myosin IB hampers the phagocytic process, mainly fail to
initiate the phagosomes. It has been hypothesized that the myosin IB interacts with the plasma membrane
and actin filament during phagosome initiation but the exact mechanism is unclear. CTD of myosin IB is a
SH3 domain, which is well known for protein-protein interaction and thus can be involved in
synchronization of signalling process and cytoskeletal activity. Thus, a structural insight into Myosin IB CTD
can be very beneficial in understanding the role of the proteins in parasite invasion process.
The structural insight into Myosin IB CTD is needed for better understanding of its interacting
partners in phagosome formation and parasite invasion process. Recombinant Myosin IB CTD was
successfully cloned and over expressed in pET21c E. coli expression vector. It was purified using Ni-NTA
chromatography followed by gel filtration chromatography. Purified protein with a concentration of 30
mg/ml was used to screen crystallization condition using commercial screens by hanging drop method.
After screening 960 conditions, only one condition from crystal Screen II (Hampton Research) in 2 M
Ammonium sulphate and 5% isopropanol at 16°C big cluster like crystals were seen obtained.
Crystals obtained were not of diffractable quality and were very temperature sensitive. Crystals
tend to dissolve after some time at microscope at room temperature. Different additives and different
temperatures were tried for crystallization. Bigger crystals were obtained at 4°C and using ethanol as an
additive but they were also in clusters. The final crystallization condition consists of 2.2 M ammonium
sulphate, 5 % isopropanol and the hanging drop consists of 1 ml of protein and precipitant each and 20%
ethanol was added only to drop at 4°C. All the processes including putting drops, observing plates, and
freezing of crystals were carried out at 4°C. Single crystals were broken from the clusters and were sent for
data collection at DBT Beamline, ESRF France. The crystals diffracted at a resolution of 1.7 Å. Structure was
solved by molecular replacement and was refined by phoenix to R-factor and Free_R factor of 19% and 21%.
CTD crystals obtained in crystal
Screen (Hampton Research)
CTD crystals after optimizing
the crystallization condition
References:
• Voigt and Guillen, Cellular Microbiology 1(3), 195-203 (1999)
• Voigt et al., Journal of Cell Science 112, 1191-1201 (1999)
• Marion et al., Cellular Microbiology 7 (10), 504–1518 (2005)
• Ravdin et al., Infect. Dis. 251, 2395–5407 (1996)
142
Poster P-68
Abstract
Crystallization of lactoperoxidase in complex with acetylsalicylic
acid, salicylhydroxamic acid and benzylhydroxamic acid
Singh AK1, Singh RP2, Sinha M2, Kaur P2, Sharma S2 and Singh TP2
1Department
2Department
of Biotechnology, Sharda University, Greater Noida, India
of Biophysics, All India Institute of Medical Sciences, New Delhi, India
Lactoperoxidase (LPO), (EC. 1.11.1.7) is a member of the family of glycosylated mammalian hemecontaining peroxidase enzymes which also includes myeloperoxiadse (MPO), eosinophil peroxidase (EPO)
and thyroid peroxidase (TPO). The binding studies of bovine lactoperoxidase with three aromatic ligands,
acetyl salicylic acid (ASA), salicylhydoxamic acid (SHA) and benzylhydroxamic acid (BHA) have indicated
that these three compounds bind to lactoperoxidase.
Co-crystallizations of LPO with ASA, SHA and BHA
The freshly purified and lyophilized samples of LPO were dissolved in 0.01 M phosphate buffer to a
concentration of 25 mg/ml. The ligands (ASA, SHA and BHA) were dissolved in the above buffer containing 30%
(v/v) methanol. Both solutions were mixed in equal volume giving protein to ligand molar ratios of approximately
1:10. A reservoir solution containing 0.2 M ammonium iodide and varying concentrations of PEG-3350 from 10%
to 30% (w/v) were prepared. 6 µl of protein-ligand solution was mixed with 6 µl of reservoir solution to prepare
12 µl of drops for hanging drop vapor diffusion method. This experiment was repeated in the three set ups of
crystallization for LPO with ASA, SHA and BHA. The rectangular and dark greenish colored crystals measuring
up to 0.3 × 0.2 × 0.2 mm3 were obtained after five days in 0.2 M ammonium iodide and 20% (w/v) PEG-3350. The
crystals obtained in that condition appeared in clusters and found to diffract poorly. Since LPO is a calcium
binding protein therefore in order to improve the crystallization process and crystal quality, 2 mM CaCl2 was
added in the above crystallization condition. This addition resulted in the growth of single crystals measuring up to
0.4 × 0.3 × 0.2 mm3. These crystals were tested in the X-ray beam and were diffracted nicely.
Initial results
After optimization
References
Singh AK et al., J. Mol. Biol. 376, 1060–1075 (2008)
Singh AK et al., Biophys. J. 96, 646–654 (2009)
Singh AK et al., J. Biol Chem. 284, 20311–20318 (2009)
143
Poster P-69
Abstract
Adapting DARPins for crystallography
Wu YF, Batyuk A, Honegger A and and Plückthun A.
Department of Biochemistry, University of Zurich, Zurich, Switzerland
Designed Ankyrin Repeat Proteins (DARPins) [1,2] are widely used as crystallization aides [3,4] and well
known for their stability, high expression yield, and their ability to crystallize in various conditions.
Moreover, DARPins can be used as a search model in molecular replacement to address the phase problem.
Despite all advantages, DARPins sometimes fail to provide the necessary polar surface for crystal contacts
due to their small size in comparison to the target protein. To extend the range of potential applications of
the DARPin technology, we have developed two new classes of DARPin based fusion proteins: (I) rigid
DARPin and -lactamase fusions (DB), (II) rigid DARPin and DARPin fusions (DD). The main purpose of the
present design was to extend the DARPin in a rigid way by another well folding protein to increase the
potential crystallization interface, and thus to extend the possibility of crystal contact formation. In the first
design, the C-terminal helix of the C-capping repeat of a DARPin extends and continues as the N-terminal
helix of its fusion partner, -lactamase (Fig 1). We designed and individually optimized six variants of the
fusion protein, which vary in the degree of overlap between the two helices and therefore in the relative
orientation of the two domains. The design of the second class was similar to the DB, by replacing the lactamase with another DARPin molecule (Fig 2). We used a similar design strategy to fuse two DARPins in
such a way that the two DARPin paratopes do not interfere with each other. The two DARPins can have the
same or different specificities. Each DARPin could bind a target protein which potentially makes crystal
contacts to increase the possibility of crystal formation for the whole complex. Nine different DAD
constructs were found to be possible that make the ligand binding sites of the two DARPins face in different
directions.
Fig. 1 DARPin-lactamase fusion. The DARPin
molecule is rigidly fused to the -lactamase via
the shared helix (experimental x-ray structure).
Fig. 2 DARPin-DARPin fusion. Two DARPin
molecule are rigidly fused head-to-tail via the
shared helix (experimental x-ray structure).
The x-ray structure of four members of DB and seven members of DD were experimental confirmed.
Moreover, two structures of each class were determined in complex with the target protein(s). The fusion
framework is generic and can be used with any DARPin, and can therefore be adapted to any target protein.
These DB and DD fusion constructs hopefully become a toolbox for robust crystal formation and phasing.
References
[1] Binz HK, et al., J.Mol. Biol., 332, 489-503 (2003)
[2] Binz HK, et al., Nature Biotechnology, 22, 575-582 (2004)
[3] Huber, T, et al., J. Struct. Biol., 159, 206-221 (2007)
[4] Sennhauser G, et al., Structure, 16, 1443-1453 (2008)
144
Poster P-70
Abstract
A strategy for selecting the pH of protein solutions to enhance
crystallization
Zhang CY1, Yin DC1,*
1Institute
for Special Environmental Biophysics, Key Laboratory for Space Bioscience and Biotechnology,
School of Life Sciences, Northwestern Polytechnical University, Xi’an, Shaanxi, People’s Republic of China
The pH of a solution is an important parameter in crystallization that needs to be controlled in order to
ensure success. The actual pH of the crystallization droplet is determined by the combined contribution of
the buffers in the screening and protein solutions, although the contribution of the latter to the pH is often
ignored. In this study, the effects of the buffer and protein solution pH values on the results of screening are
systematically investigated. It was found that varying the protein solution pH can result in a low
crystallization success rate. To rationally utilize the pH of protein solutions for improved crystallization, we
propose the following strategy for selecting appropriate protein solution pH values without any initial
testing: (i) when screening with only one protein solution, the pH should be as low, as high or as divergent
from the pI as possible for a basic, acidic or neutral protein, respectively, within its stable pH range; (ii)
when screening with two protein solutions, the pH values should be well separated from one another; and
(iii) when multiple pH values are utilized, an even distribution of pH values is the best approach to increase
the success rate of crystallization (also applicable for biochemically uncharacterized proteins).
Figure 1 The number of non-redundant crystallization hits against the number of protein solution pH values
for two buffer systems. A-F represents lysozyme, concanavalin, chymotrypsinogen A, catalase, glucose
isomerase, and myoglobin, respectively. 1 and 2 represent single-component buffer and multiplecomponent buffer system, respectively. The upper and lower limits are not error bars but rather represent
the maximum and minimum number of crystallization hits, respectively. The black squares between the
upper and lower limits are the average number of hits for the specific number of protein solution pH
combinations and the red triangles represent the number of hits using the protein solution pH selection
strategy proposed herein.
Acknowledgements
This work was supported by the National Natural Science Foundation of China (Grant Nos. 31170816,
11202167, 51201137, 31200551), and the China Postdoctoral Science Foundation (Grant No. 2013T60890),
the Natural Science Basic Research Plan in Shaanxi Province of China (Grant Nos. 2012JQ3009, 2013JQ4032,
2014JM4171).
References
Zhang CY, et al., Acta Crystallogr. Sect. F Struct. Biol. Cryst. Commun. F69(7), 821–6. (2013)
Newman, J. Acta Cryst. D60, 610–2. (2004)
Dierks K, et al., Acta Crystallogr. F Struct. Biol. Cryst. Commun. 66, 478–84. (2010)
Kantardjieff KA & Rupp B, Bioinformatics 20, 2162–8. (2004)
145
Poster P-71
Abstract
Towards investigation of structural dynamics of rhodopsin with
XFEL
Wu W, Nogly P, Rheinberger J, Tsai CJ, Standfuss J, Li XD, Deupi X, Panneels V,
Schertler G
Laboratory of Biomolecular Research, Paul Scherrer Institute, Villigen, Switzerland
Rhodopsin is a retinal-binding protein responsible for dim light vision and is a prototypic
G-protein coupled receptor (GPCR). The chromophore 11-cis-retinal is associated with the
protein via a protonated Schiff base linkage to K296. Upon light activation, the
photoisomerization of the chromophore and local rearrangements happen in
femtoseconds to picoseconds followed by helical movements in milliseconds (Schertler,
2005). The dark-state structure of rhodopsin has been solved (Edwards et al, 2004; Okada
et al, 2004) but much less is known about other intermediates in particular the early ones.
The highly intense and femtosecond pulses of the X-ray free electron laser (X-FEL) will
allow detection of ultrafast changes (Aquila et al, 2012). Time-resolved femtosecond
crystallography (TR-SFX) and wide-angle x-ray scattering (TR-WAXS) will be used in
parallel to unravel the structural basis of the rhodopsin photocycle.
References
Schertler G, Curr. Op. Struct. Biol. 15, 408-415 (2005)
Edwards P et al, J. Mol. Biol. 343, 1439-1450 (2004)
Okada T et al, J. Mol. Biol. 342, 571-583 (2004)
Aquila A et al, Opt Express. 20, 2706-2716 (2012)
146
Poster P-72
Abstract
From the frying pan into the fire: overcoming cryo-cooling induced
twinning by room temperature data collection using the HC1 at
ESRF beamline ID29
Monteiro DCF1, Bravo JP2, Patel V1, Webb ME1, Pearson AR2
1.
2.
Astbury Centre for Structural Molecular Biology, School of Chemistry, University of
Leeds, Leeds, UK
Hamburg Centre for Ultrafast Imaging, Institute of Experimental Physics, University of
Hamburg, Hamburg, Germany
E. coli Aspartate α-decarboxylase undergoes a post-translational backbone rearrangement
and cleavage to generate the pyruvoyl cofactor required for catalysis. This posttranslational modification is catalysed by the recently identified processing factor PanZ. We
obtained crystals of the ADC4.PanZ4 complex in a variety of conditions that diffracted to at
best 2.9 Å but all attempts at structure solution were hampered by indexing ambiguities
and twinning. Packing analysis based on the cell dimensions suggested multiple copies of
the oligomer in the asymmetric unit. In an attempt to improve the diffraction quality of the
crystals, we planned a room temperature dehumidification experiment at the ESRF using
the humidified gas stream HC1 on beamline ID29. To our surprise, the crystals diffracted to
beyond 1.5 Å with no twinning. We will present the overall complex structure, as well as
details of further crystal optimisation for ongoing mechanistic studies.
147
Poster P-73
Abstract
Crystal systems for structure-based drug design
Breed J
AstraZeneca, UK
The aim of this presentation is to describe and illustrate the idea of a crystal system (a set
of reagents and activities that can reliably be used over and over again to produce
structures of target proteins with a wide variety of ligands) and the key parameters that
make such a system suitable for iterative structure-based drug design. The presentation
will also emphasize the utility of crystal systems in industrial and academic settings and
encourage all present to develop crystal systems to generate leads for drug design.
148
Poster P-74
Abstract
Influence of precipitants on the thermodynamic parameters of
protein crystallisation
Stavros P1, Nounesis G1, Saridakis E2
1Biomolecular
Physics Laboratory, INRASTES, National Centre for Scientific Research “Demokritos”, Athens, Greece;
of Structural and Supramolecular Chemistry, INN, National Centre for Scientific Research “Demokritos”,
Athens, Greece
2Laboratory
In crystallisation trials, different chemical environments will lead to very different and a priori unpredicatable
outcomes. Supersaturation attained and the specific crystal contacts that may be formed by the intermediary of solution
components cannot provide a full explanation of the radically varying crystallisation behaviour induced in different
protein-buffer systems by different precipitating agents. Thermodynamic parameters and the way in which they are
influenced by various components of the system can provide additional understanding of the crystallisation process.
When molecules previously in solution form a crystalline phase, their entropy is reduced. The crystallization driving
force must then come either from the enthalpy of the intermolecular interactions in the crystal (ΔHcryst), or from a gain
of entropy of the solvent (TΔSsolv). There is indeed often an entropy gain upon crystallisation, since a large proportion of
the water molecules previously forming ordered hydration shells around the macromolecules are excluded from the
crystal (Vekilov et al., 2002). Enthalpic gain through formation of crystal contacts is the other crystallisation driving
force.
The importance of both entropic and enthalpic effects for crystallisation suggests that changes in these parameters in
the solution, brought about by the precipitant and buffer, will be crucial to the outcome. One possible way to think about
this is that a system at low entropy that can gain much entropy by yielding crystals (for example one with well-ordered
solvent due to extended and deep hydration shells around the protein molecules) would provide an ideal driving force
for crystallisation. A system that is at high entropy from the outset would not be entropically favourable to
crystallisation, since the driving ΔS is likely to remain small. The enthalpic effect should also be taken into account: in
cases where the enthalpy of crystallization is very favourable it will be the main driving force, whereas if the enthalpic
contribution is small or even unfavourable, even a small solvent entropic effect may become crucial.
To identify such effects, we have used Differential Scanning Calorimetry in order to gather data regarding the stability at
crystallisation temperatures of protein molecules, in mixtures that are known to promote crystallisation versus
mixtures where the given protein does not crystallise. Both the ΔG (Free Energy) of unfolding at different temperatures
and salt concentrations and its breakdown into entropic and enthalpic components were obtained. Results for lysozyme
indicated that in sodium acetate buffer at pH 4.5 and in increasing concentrations of salts in which it crystallises,
lysozyme is enthalpically stabilised (inceasingly positive ΔH of unfolding) and entropically destabilised (increasingly
positive ΔS of unfolding), with an overall stabilising ΔG. The opposite effects were observed for salts in which lysozyme
does not crystallise. Preliminary results for RNase A indicate a similar trend.
Concentrating on entropic effects, one possible explanation would be the following. A greater ΔS of unfolding can be the
result of either a reduction of the entropy of the native state or an increase in the entropy of the unfolded state. In the
former case, we have a lower entropy of the native state in solution and therefore, according to the above argument, a
greater entropic driving force for crystallisation. These findings suggest the possibility of using such measurements for
a priori precipitant selection.
Thermodynamic stability curves for lysozyme in sodium
acetate buffer at pH 4.5 and various concentrations of
Na2SO4, a salt in which it crystallises.
Reference
Vekilov PG et al., Acta Cryst. D58, 1611-1616 (2002)
149
Poster P-75
Abstract
Crystallization of proteins on functionalized surfaces indicate a
non-classical mechanism for the heterogeneous nucleation
Fermani S.1, Falini G.1, Vettraino C.1, Gavira JA2,and Garcia Ruiz JM2
1Department
of Chemistry “G. Ciamician”, University of Bologna, Bologna, Italy; 2Laboratorio de Estudios
Crystalograficos, Instituto Andaluz de Ciencias de la Tierra (CSIC-UGR), Granada, Spain.
The study of the three-dimensional crystallographic structures of proteins is a starting point to
understanding their structure–function relationships. This usually requires single crystals of high diffraction
quality. Since there is no well defined law that allows the determination of which chemical and physical
experimental conditions can lead a protein to crystallize, a ‘trial-and-error’ approach is usually applied. The
studies conducted on protein crystallization generally focus on how to maximize the probability of
nucleation, i.e. the nucleation frequency (Garcia-Ruíz, JM, 2003), this being the critical step of the whole
crystallization process, by introducing in the system any material that decreases the energy nucleation
barrier, facilitating the crystal formation at lower supersaturation values (Nanev, CN, 2007). Functionalized
mica sheets and polystyrene films exposing ionisable groups, amino or sulfonated groups respectively, and
with different densities of charged groups have been used as heterogeneous nucleating surfaces for model
proteins. Crystallization trials were carried out using the vapour diffusion method. The results show that
using these surfaces, the starting protein concentration necessary to form crystals is reduced and a decrease
of the waiting time (an estimation of the nucleation time) was also observed for some proteins, suggesting a
surface stabilization effect on crystal nuclei (Tosi G, et al., 2008). Moreover the results show that (i) the
heterogenenous nucleation does not crucially affect the nucleation process under conditions of high
supersaturation and that (ii) the nucleating action of the functionalized surfaces is mainly related to their
superficial charge distribution more than to their absolute charge, according to both classical and colloidal
nucleation theories (Tosi G, et al., 2011). The molecular mechanism by which heterogeneous agents
promote the nucleation is still unclear, although examples of epitaxial crystallization have been reported. A
focused set crystallization trials using sulphonated polystyrene films point out that the process of
heterogeneous nucleation can be described by a non-classical mechanism. We propose that due to
concentration fluctuations, unstable pre-nucleation aggregates homogeneously form in solution in
conditions of low supersaturation, when the chemistry of solution favours this event (Fermani S, et al.,
2013). Then, heterogeneous substrates stabilize pre-nucleation clusters by non-specific interactions and
trigger the nucleation event. This concept applied to proteins, but also to any system where the ordered
assembly is governed by weak interactions, highlights the importance of the chemistry of the solution as a
key parameter to achieve nucleation, even in the presence of heterogeneous nucleants.
Cartoon representation of the two-steps mechanism proposed for the
heterogenous nucleation of proteins at low supersaturation. Prenucleation clusters form in solution because of the fluctuations of
protein concentration, but the probability that they evolve in a crystal
nuclei is quite low (red and angry face). The heterogenous nucleant
surface (AFM image) stabilize the clusters formed in its proximity
(yellow and serious face) drastically increasing their chance to convert
in crystalline nuclei (green and happy face).
References
Garca-Ruiz, J M, J. Struct. Biol., 142, 22–31 (2003).
Nanev, CN, Cryst. Res. Technol., 42, 4–12 (2007).
Tosi G, et al., Acta Crystallog D64, 1054-1061 (2008).
Tosi G, et al., Cryst. Growth Des. 11, 1542–1548 (2011).
Fermani S, et al. Cryst. Growth Des., 13, 3110−3115 (2013).
150
Poster P-76
Abstract
Microfluidics for in situ following mineralization at organic
surfaces by Micro-Raman
Liszka B1, Rho HS3, Yang YS1, Lenferink ATM1, Witkamp G-J2, Terstappen L1,Otto C1
1 Department
of Medical Cell BioPhysics, MIRA institute, University of Twente Enschede, The Netherlands
University of Technology Process & Energy Laboratory, Delft, The Netherlands
3Department of Mesoscale Chemical System, MESA institute University of Twente Enschede, The
Netherlands)
2 Delft
We present a method for in situ detection of sub-micron to nano-sized crystals formation on various
nanometer thin organic membranes by micro Raman spectrometry and imaging.
Mineralization has been assumed to be highly dependent on the nature of surface. A number of polymer
materials was selected for mineralization experiments, for example: polysulfone, poly(p-phenylene-oxide),
polystyrene . In order to observe the smallest possible nano-crystals at a polymer surface the Raman
scattering of the crystals must be of the some order of magnitude as the Raman scattering from the organic
layer. It was therefore decided to prepare very thin polymer films with a thickness of the order of
nanometers as part of the microfluidic device.
The microfluidics system has been optically coupled with a micro Raman spectrometer in order to follow
in situ crystal growth at an organic membrane interface with the ionic solution. The chip has been design as
an inverted system with fluidic channels positioned on top and covered with organic film on CaF2 disk. The
advantage of this design is that it enables acquisition of high resolution images and detection of nanoobjects. Moreover, a microfluidic system facilitates control over mass transport, such that experiments can
be carried out in a confined volume and under well-defined conditions in solution. Additionally, Ramanbased micro-spectroscopic methods like spontaneous Raman scattering and coherent Raman imaging can
be applied on samples under ambient conditions in the micro-fluidic device in real time. Moreover, the
Raman spectra of minerals are directly informative with respect to the chemical composition and can be
directly interpreted in terms of the symmetry of the crystals in case of different polymorphs. Furthermore, a
great advantage is the sensitivity which is sufficient to detect nano-sized mineral particles
Finally, a model super saturated mineralization solution was prepared to understand early phenomena
occurring in solution. The solution was investigated by a double pulse experiment, which enables to assess
the mineralization rate and gives insight in the nature of pre-nucleation prior to a mineralization process. A
model mineralization solution has a well-defined metastable region which depends on parameters, such as:
supersaturation ratio, pH , ionic strength, temperature. The solution was applied in microfluidics to
investigate crystals growth in situ at organic membrane.
151
Poster P-77
Abstract
Protein crystallization with microseed matrix screening:
application to human germline antibody Fabs
Obmolova G, Malia TM, Teplyakov A, Sweet RW, Gilliland G
Janssen Research & Development LLC, Spring House, Pennsylvania, USA
The crystallization of 16 human antibody Fab fragments constructed by combining four
different heavy chains and four different light chains was enabled by employing microseed
matrix screening (MMS). In initial screening (288 conditions), diffraction quality crystals
were obtained for only 3 Fabs, while many Fabs produced hits that required optimization.
Application of MMS, using the initial screens and/or refinement screens, resulted in
diffraction quality crystals for these Fabs. Five Fabs that failed to give hits in the initial
screen were crystallized by cross-seeding MMS followed by MMS optimization. The
crystallization protocols and strategies that resulted in structure determinations of all 16
Fabs are presented. These results illustrate the power of MMS and provide a basis for
developing future strategies for macromolecular crystallization.
152
Poster P-78
Abstract
Cryoprotection and ligand solubilization: Evaluation of
multicomponent mixtures
Ciccone L1,2, Vera L1, Tepshi L1,2 and Stura EA1
1CEA,
iBiTec-S, Service d’Ingénierie Moléculaire des Protéines, Laboratoire de Toxinologie Moléculaire et
Biotechnologies, Gif-sur-Yvette, France; 2Dipartimento di Scienze Farmaceutiche, Università di Pisa, Pisa,
Italy.
Crystallographic structure determination of protein-ligand complexes is an important step in the progress
from a low affinity lead compound to an active molecule. The combination of low affinity and low solubility is
a hurdle for structural studies of ligand-protein complexes. Complete solubilization of hydrophobic ligands is
required for their use in co-crystallization or crystal soaking experiments to obtain interpretable electron
density maps for the ligand. High-throughput screening is used to select among a library of millions of
compounds those that bind to a target protein, inhibit a particular enzymatic reaction or block a cellular
transport mechanism. Typically the chemical compounds are dissolved in dimethyl sulfoxide (DMSO)[1] to
produce an aqueous solution. This solution is diluted during the screening to ensure that the concentration of
DMSO does not exceed 10%; as higher concentrations can be damaging to the target protein.[2] The solubility
of ligands is a problem for the crystallization of protein-complexes, more so when the ligand or fragment has
a low affinity for its target. For co-crystallization the ligand must be soluble in the crystallization precipitant
so as to be in excess compared to the protein. In a previous study, by mixing cryo-preserving compounds that
act as precipitants with compounds that have the opposite effect we developed a set of multicomponent
mixtures that could be combined with a precipitant and a buffer so as to be able to prepare crystals for X-ray
data collection at high intensity synchrotron facilities without hassle.[3] These mixtures can stabilize crystals
for periods long enough for ligand soaking experiments. We report an extended set of multicomponent
solutions for crystal cryoprotection which includes additional components, namely dioxane and butanediol
and analyse the use of dioxane for ligand solubilization, alone and in conjunction with other cryoprotectant
components and its compatibility for protein crystallization and crystal soaking. Dioxane has been
extensively used as an additive in macromolecular crystallization for its ability to mediate lattice
interactions.[4] Ligand insolubility is also a common problem in chemistry and a lot of compounds are
insoluble in organic solvents. There is no general rule on how to dissolve chemical molecules,[5] and the wellknow phrase "similia similibus solvuntur" (polar solvents dissolve polar compounds and non-polar solvents
dissolve nonpolar compounds best) is helpful only as a general guide Figure 1.
The strategy of using a mixture of solvents to solubilize ligands
is well known. Two or more solvents together enhance the
solubility of insoluble compounds.[6]This approach of mixing
together compounds was used to create solutions that are
effective in solubilizing hydrophobic ligands for cocrystallization and soaking experiments. Since soaking
experiments are best done during the cryoprotection step,
these solutions must be formulated to also inhibit ice
formation. We report the co-crystallization tests using the
model protein transthyretin (TTR) with various hydrophobic
ligands solubilized using these solvent mixtures. High
resolution data was collected from crystals cryo-protected in
solutions formulated by mixing the crystallization precipitant,
Figure 1: Classification of solvent selectivity
a buffer and these mixed solubilization compounds.
[1] Fedarovich A, et al., PloS One 7, e44918 (2012)
[2] Jackson M, Mantsch HH. Biochimica et Biophysica Acta 1078, 231–235 (1991)
[3] Vera L, Stura E A. Crystal Growth & Design 14, 427–435 (2014)
[4] Ménétrey J, et al., Crystal Growth Design 7, 2140–2146 (2007)
[5] Barton AFM, Chemical Reviews 75, 731–753 (1975)
[6] Jouyban A, Journal of Pharmacy & Pharmaceutical 11, 32–58 (2008)
153
Poster P-79
Abstract
Calcium concentration is crucial for crystallization of calcium
binding protein-5 from Entamoeba histolytica
Kumar S1 and Gourinath S1*
1School
of Life Sciences, Jawaharlal Nehru University, New Delhi, Delhi, 110067, India
Entamoeba histolytica is the causative agent of human amebiasis. Phagocytosis is the
major route of this parasite’s food intake and is responsible for its virulence. Calcium and
calcium binding proteins play major roles in its phagocytosis. The calcium binding
protein-5 from E. histolytica (EhCaBP5) is a cytoplasmic protein; its expression is very
sensitive to serum starvation and seems to be involved in binding with myosin IB. In this
study, EhCaBP5 was cloned, expressed in Escherichia coli, and purified by affinity and sizeexclusion chromatography. While gel filtration chromatography elution buffer contained
50 mM Tris pH 8.0, 30 mM NaCl, 0.2 mM CaCl2, 5 mM -mercaptoethanol and eluted peak
fraction were concentrated to 10 mg/ml. Initial crystallization trials were carried out by
the hanging drop vapour-diffusion method in 96-well plates using a crystallization robot.
200 nL protein solution was mixed with an equal volume of precipitant solution and
equilibrated against 100 ml reservoir solution (precipitant). The plates were incubated at
289 K. After 10–15 days of equilibration, tiny crystals or needles appeared in Index
condition No. 24 (2.8 M sodium acetate pH 7.0) and 72 [0.2 M MgCl2, 0.1 M bis-tris pH 5.5,
25%(w/v) PEG 3350], Crystal Screen condition No. 23 [0.2 M MgCl2,0.1 M Na-HEPES pH
7.5, 30%(v/v) PEG 400], SaltRx condition No. 2 (0.1 M bis-tris propane pH 7.0, 2.8 M
sodium acetate pH 7.0) and Natrix Screen condition No. 1 (0.01 M MgCl2, 0.05 M MES pH
5.6, 1.8 M lithium sulfate), all from Hampton Research (Aliso Viejo, California, USA). The
conditions were further optimized in 24-well plates by varying pH values in the range 5.0–
7.0 and concentrations of sodium acetate, lithium sulfate and PEG. Surprisingly at high
concentrations of CaCl2 (>1 mM) protein precipitated in the crystallization drop and we
never get crystals in high CaCl2 concentrations. Diffraction quality crystals were obtained
in 2.6 M-2.8 M sodium acetate, 0.1 M bis-tris pH 5.5-5.8 and no additional CaCl2 was added
in the crystallization condition. Finally, we have determined the crystal structure of
EhCaBP5 at 1.9 Å resolution in the Ca2+-bound state, in which EF hand motif shows an
unconventional mode of Ca2+ binding coordination to a canonical EF-hand motif.
154
Poster P-80
Abstract
Trace particles everywhere, in situ DLS shows its adaptability
Meyer A1, Falke S2, Dierks K1, Schubert R2, Betzel C2
1XtalConcepts GmbH,
Marlowring 19, 22525 Hamburg, Germany
2Institute
of Biochemistry and Molecular Biology, Laboratory for Structural Biology of
Infection and Inflammation, University of Hamburg, c/o DESY, Build. 22a, Notkestr. 85,
22603 Hamburg, Germany
Particle detection based on dynamic light scattering (DLS) is well known, widely used.
However, DLS is technically realized mostly in standard laboratory desktop instruments.
Despite the fact that those instruments are designed to apply DLS in cuvettes, this method
is far more versatile and adaptable as commonly expected. So, DLS could be applied on
many light transparent containers such, crystallization plates [1], as capillaries [2], microchannel plates, dialysis chambers and even ampules, flasks and experimental containers.
In course of that, we named this method in situ DLS. DLS measurements in such different
vessels, required the development of a small external DLS optical head, containing all
required optical elements. Once finalization of this optical unit has been archived, it was
adaptable on almost anything that was transparent for visible light. Some extensions and
variations of that unit have been developed too, e. g. a multi-channel DLS, that analyzes
different measurement volumes along a laser beam and evaluates them independently.
Developments of desktop instruments based on in situ DLS have been archived as well, for
example XtalController 900 [3-4] and SpectroLight 600. Here we present some
remarkable examples that we could arrange in cooperation with the University of
Hamburg and the European Molecular Biology laboratory (EMBL).
References
[1] Dierks K et al., Crystal Growth and Design, No. 8, 1628 (2008)
[2] Oberthuer D et al., PLoS One 7(6), p. e33545 (2012)
[3] Meyer A et al., Acta Crystallogr Sect F Struct Biol Cryst Commun 68(Pt 8), 994-8 (2012)
[4] Vekilov PG, Nanoscale 2, 2346-7 (2010)
155
Abstract
Poster P-81
Mitochondrial peroxidase from Leishmania integrates signals
from calcium and reactive oxygen species
Morais MAB1, Giuseppe PO1, Souza TACB2, Honorato RV1, Alegria, T3, Oliveira PSL1,
Oliveira M4, Tomas AM5, Netto LES3, Murakami MT1
1Laboratório
Nacional de Biociências, Centro Nacional de Pesquisa em Energia e Materiais,
Campinas/SP, Brazil; 2Laboratório de Proteômica e Engenharia de Proteínas, Instituto Carlos Chagas,
Fiocruz, Curitiba/PR, Brazil; 3Departamento de Genética e Biologia Evolutiva, Instituto de Biociências,
Universidade de São Paulo, São Paulo/SP, Brazil; 4Departamento de Biologia, Universidade Estadual
Paulista Júlio de Mesquita Filho, Campus do Litoral Paulista São Vicente, São Paulo/SP, Brazil; 5Instituto
de Biologia Molecular e Celular, Universidade do Porto, Porto, Portugal.
The mitochondrial tryparedoxin peroxidases (mTXNPxs) participate in regulatory pathways involving
reactive oxygen species (ROS) and play an important role in the virulence of trypanosomatids. Once upregulated, they protect these parasites from H2O2-induced apoptosis, a process controlled by the
interplay among ROS generation, Ca2+ overload and mitochondrial dysfunction. As typical member of
the 2-Cys peroxiredoxins family, the catalytic cycle of mTXNPx includes an oxi-reduction process and
shift in enzyme quaternary structure (dimers and doughnut oligomers). The overall goal of this work is
shed light on the molecular mechanisms involved in the catalytic and oligomeric cycles in TXNPx. The
mitochondrial TXNPx of Leishmania braziliensis (mLbTXNPx) was cloned and overexpressed in
prokaryotic system, purified and crystallized. We solved the crystal structures of oxidized mLbTXNPx
in both dimeric and decameric forms. Different techiniques such as size-exclusion chromatography,
dynamic light scattering and Small-angle X-ray scattering were used to study the oligomerization of
mLbTXNPx. Peroxidasic assays, site-directed mutagenesis and molecular dynamics simulations were
also performed. Our analyses indicate that protein decamerizes upon calcium binding, which is not
observed for other Prxs and that Ca2+ enhances the peroxidase activity of mLbTXNPx by stabilizing the
dimer-dimer interface. The crystal structures of mLbTXNPx along with mutagenesis and biochemical
studies revealed the molecular determinants for Ca2+ regulation, the mechanism involved in the
pH-dependent dimer-to-decamer transition and provided insights into the catalytic cycle of AhpC/Prx1
subfamily members. Together, our findings point out the calcium-dependent decamerization as a
unique feature of leishmanial mTXNPxs, unveiling a potential parasite-specific crosstalk between Ca2+
and ROS signaling pathways via mTXNPxs in the Leishmania giant mitochondrion.
Keywords: Peroxiredoxin, structure, oligomerization.
Acknowledgements: FAPESP and CAPES.
References
Wilkinson SR et al., J Biol Chem, 278(34), 31640-6 (2003)
Barr SD and Gedamu L, J Biol Chem, 278(12), 10816-23 (2003)
Iyer JP et al., Mol Microbiol, 68(2), 372-91 (2008)
Piacenza L et al., Biochem J, 410(2), 359-68 (2008)
Alvarez MN et al., J Biol Chem, 286(8), 6627-40 (2011)
156
Abstract
Poster P-
157
Abstract
Poster P-
158
Notes
159
Notes
160
Notes
161
Notes
162
List of presentation authors (in alphabetical order)
name
no.
page
name
no.
page
Aagaard
Abdallah
A
BG
P-32
O-18
107
47
Biscans
Blagova
B
EB
O-20
P-45
49
120
Abdallah
BG
O-43
72
Blankenship
RE
P-23
98
Abdulrehman
SA
P-67
142
Bogan
M
O-28
57
Abrahams
JP
O-15
44
Bonneté
F
O-20
49
Adachi
H
O-35
64
Bonneté
F
O-5
34
Adam
V
P-31
106
Böszörmenyi
A
P-13
88
Adawy
A
O-18
47
Boudes
M
O-21
50
Aizel
K
O-21
50
Bourenkov
G
O-23
52
Akella
S
O-31
60
Bourgeois
D
P-31
106
Amberg
C
P-28
103
Boutet
S
O-28
57
Andersson
KK
P-55
130
Boyko
KM
P-19
94
Appio
R
P-64
139
Boyko
KM
P-20
95
Aquila
A
O-25
54
Brandt van Driel
T
O-28
57
Aquila
A
O-28
57
Bravo
JP
P-72
147
Arnlund
D
O-28
57
Breed
J
P-73
148
Aschauer
P
P-17
92
Breinbauer
R
P-17
92
Baba
S
P-12
87
Brennich
ME
O-20
49
Bäckström
S
P-32
107
Brezovsky
J
P-22
97
Balaev
VV
P-60
135
Broadley
SG
P-38
113
Barcena
J
P-66
141
Brown
M
P-4
79
Barco
J
P-35
110
Bruno
AE
O-11
40
Bari
S
O-28
57
Budayova-Spano
M
P-54
129
Barnes
C
O-27
56
Busse
RA
P-9
84
Barret
L-A
O-5
34
Byrdin
J
P-31
106
Barty
A
O-28
57
Caffrey
M
O-40
69
Basu
S
P-4
79
Calero
M
O-27
56
Batyuk
A
P-69
144
Calero
G
O-27
56
Benz
J
O-6
35
Cao
HL
O-37
66
Bergoin
M
O-21
50
Cao
HL
P-65
140
Bernacchioni
C
P-25
100
Casaite
V
P-15
90
Berntsen
P
O-28
57
Castro
F
P-29
104
Berzin
C
P-54
129
Cerk
IK
P-13
88
Betzel
C
P-21
96
Chaloupkova
R
P-22
97
Betzel
C
P-80
155
Chaloupkova
R
P-26
101
Betzel
C
P-57
132
Chang
WC
P-28
103
Betzel
C
P-60
135
Chapman
HN
O-23
52
Betzel
C
P-62
137
Chapman
HN
O-28
57
Bird
L
P-13
88
Chawla
U
P-4
79
Birner-Grünberger
R
P-17
92
Chen
Z
P-36
111
163
name
no.
page
name
no.
page
Doak
Dohnálek
RB
J
O-28
P-14
57
89
J
P-49
124
Dreveny
I
P-17
92
Chen
Cherezov
RQ
V
P-65
O-2
140
31
Chum
KKC
P-28
103
Ciccone
L
P-78
153
Clabbers
M
O-15
44
Dreyer
M
P-51
126
Cochrane
G
P-44
119
D
P-14
89
Coe
J
P-4
79
P-49
124
A
O-27
56
M
P-13
88
Colletier
J-P
P-31
106
Dušková
J
Cohen
Dušková
Edman
K
P-32
107
Conejero-Muriel
M
P-48
123
Edwards
B
P-35
110
Conrad
C
P-4
79
Eriksson
P-O
O-7
36
Conrad
C
P-6
81
Fabis
R
P-51
126
Cordeiro
RSC
P-8
83
Falini
G
P-75
150
Correia
MAS P-10
85
Falke
S
P-80
155
Correia
JDG
85
Faraldo-Gómez
JD
P-7
82
Correia
MAS P-8
83
Fazio
VJ
O-39
68
Coulibaly
F
O-21
50
K
P-14
89
Dada
O
P-36
111
P-49
124
M
P-15
90
Fermani
S
P-75
150
Dahani
M
O-5
34
Fejfarová
K
Dagys
Fejfarová
Ferreira
A
P-29
104
Damas
AM
P-5
80
Ferreira
C
P-5
80
Damborsky
J
P-22
97
Ferrer
JL
P-54
129
Damborsky
J
P-26
101
Fisher
SZ
O-32
61
Daniel
L
P-22
97
Fisher
M
P-37
112
Das
KMP P-13
88
Forsyth
T
O-12
41
Datwani
S
P-35
110
Fraden
S
O-31
60
Davidsson
J
O-28
57
Frank
M
O-28
57
Davies
GJ
P-45
120
Fredslund
F
P-1
76
de Grip
W
O-18
47
Fredslund
F
P-64
139
Degtjarik
O
P-26
101
Friis
EP
P-45
120
Demeshkina
N
P-16
91
Fromme
P
P-4
79
Demidkina
TV
P-20
95
Fromme
P
P-6
81
Deng
Y
P-50
125
Fromme
P
O-18
47
DePonte
D
O-27
56
Fromme
P
O-26
55
DePonte
DP
O-28
57
Fromme
R
O-28
57
Deupi
X
P-71
146
Fromme
P
O-28
57
Di Pisa
F
P-25
100
Fromme
P
O-43
72
Dierks
K
P-80
155
Fu
X
O-27
56
Dierks
K
P-57
132
Fufina
TY
P-52
127
Dittmann
E
P-18
93
Fujimoto
M
P-40
115
P-10
Dohnálek
Eder
164
name
no.
page
name
no.
page
Fukumoto
Fushinobu
N
S
P-56
O-33
131
62
Håkansson
Hallberg
M
E
P-64
P-53
139
128
Gabdoulkhakov
AG
P-52
127
Hammerstad
M
P-55
130
Gabdoulkhakov
AG
P-60
135
Hargreaves
D
P-42
117
Gabdoulkhakov
AG
P-62
137
Harris
D
P-35
110
Garcia Ruiz
JM
P-75
150
Hašek
J
P-49
124
Garriga
D
O-21
50
He
J
O-37
66
Gautam
G
P-67
142
He
JH
O-37
66
Gavira
JA
P-48
123
Heng
JYY
P-28
103
Gavira
JA
P-75
150
Henning
R
O-28
57
Genderen
E
O-15
44
Herrmann
C
P-11
86
Gerwert
K
P-11
86
Hersleth
H-P
P-55
130
Ghini
V
P-25
100
Hessler
G
P-51
126
Giegé
R
O-24
53
Heymann
M
O-31
60
Giglio
M
O-17
46
Hiess
A
O-32
61
Gilliland
G
P-77
152
Hijnen
M
O-21
50
Goldman
A
P-58
133
Hinniger
A
P-43
118
Gorbacheva
MA
P-19
94
Hiraki
M
P-39
114
Gorrec
F
O-9
38
Ho
TCT
P-28
103
Gossert
A
P-43
118
D
P-34
109
Gourinath
S
P-67
142
R
P-41
116
Gourinath
S
P-79
154
Hocková
Hofmann
E
P-11
86
Grant
TD
O-11
40
Hofmann
E
P-23
98
Groenhof
G
O-28
57
Honegger
A
P-69
144
Grönberg
G
O-7
36
Honnappa
S
P-43
118
Grotjohann
I
O-28
57
Hou
H
P-63
138
Gruber
K
P-17
92
Huang
LJ
O-37
66
Gruner
SM
O-31
60
Hunt
JF
O-11
40
Gualtieri
EG
P-37
112
Hunter
MS
O-28
57
Guddat
LW
P-34
109
Iermak
I
P-26
101
Guder
S
P-61
136
Igarashi
K
O-33
62
Gudim
I
P-55
130
Iizuka
I
P-40
115
Guha
S
O-30
59
Ikeda-Ohtsubo
W
P-22
97
Guillet
P
O-20
49
Inaka
K
P-40
115
Guillon
V
P-31
106
Inoue
T
O-35
64
Guo
YZ
O-37
66
Ishida
T
O-33
62
Guo
WH
O-37
66
Izaac
A
P-43
118
Gutmann
S
P-43
118
Jakobi
AJ
O-23
52
Gutowski
S
P-36
111
James
D
O-28
57
Hackenberg
C
P-18
93
James
D
P-6
81
Hoffmann
165
name
no.
page
name
no.
page
Jenkins
Jiang
J
J
P-44
P-23
119
98
Kosheleva
Kotlov
I
MI
O-28
P-20
57
95
Johansson
LC
O-28
57
Kötting
C
P-11
86
Jonasson
E
O-7
36
Koudelakova
T
P-22
97
Juárez-Santiesteban H
P-24
99
Koval
T
P-49
124
Jungas
C
O-5
34
Kovaleva
E
O-27
56
Junius
N
P-54
129
Krasnoselska
OG
P-7
82
Jüttner
P
O-34
63
Krizsan
A
P-41
116
Kajander
T
P-58
133
Kubicek
J
P-51
126
Kamashev
DE
P-19
94
Kühlbrandt
W
O-38
61
Kamiya
N
P-12
87
Kühnel
K
P-9
84
Kapis
S
P-21
96
Kulathila
R
P-30
105
Karr
LJ
P-3
78
Kulikova
VV
P-20
95
Kasai
Y
P-33
108
Kumar
S
P-79
154
Kato
R
P-39
114
Kumasaka
T
P-12
87
Katona
G
O-28
57
Künz
M
P-57
132
Kaur
P
P-68
143
Kupitz
C
P-4
79
Kawano
Y
P-12
87
Kupitz
C
P-6
81
Kawasaki
M
P-39
114
Kupitz
C
O-18
47
Kenis
PJA
O-30
59
Kupitz
C
O-28
57
Keough
DT
P-34
109
Kupitz
Ch
O-43
72
Kestilä
VP
P-58
133
Kuriyama
M
P-56
131
Kimani
S
P-2
77
Kuroki
K
P-33
108
Kimbung
R
P-64
139
Kusaka
K
O-33
62
Kirian
RA
O-28
57
Kusaka
H
P-33
108
Kirkwood
JS
P-42
117
Kuta Smatanova
I
P-26
101
Kita
Kjær
S
P-33
108
Kuta Smatanova
I
P-22
97
KS
O-28
57
Kuta Smatanova
I
P-47
122
Klyszejko
AL
P-7
82
Kuty
M
P-22
97
Knoops
K
O-23
52
Kuty
M
P-26
101
Kobayashi
M
P-56
131
Laatsch
B
O-34
63
Kobe
B
O-41
70
Lalli
D
P-25
100
Koch
E
P-43
118
Lamzin
V
P-18
93
Kock
K
P-11
86
Langer
T
P-51
126
Kogan
K
P-58
133
Lashkov
AA
P-60
135
Kolenko
P
P-14
89
Lashkov
AA
P-62
137
Kolenko
P
P-49
124
Leisico
F
P-10
85
Komadina
D
O-23
52
Lenferink
ATM P-76
151
Korgenevsky
DA
P-19
94
Leone
V
P-7
82
Korgenevsky
DA
P-20
95
Li
Y-W
O-15
44
166
name
no.
page
name
no.
page
Li
Li
J
J
O-37
P-50
66
125
Mayer
Mayrhofer
N
J
P-17
P-23
92
98
Li
XD
P-71
146
Meier
T
P-7
82
Liang
M
O-23
52
Meskys
R
P-15
90
Liang
M
O-28
57
Messerschmidt
M
O-23
52
Lin
G
O-27
56
Messerschmidt
M
O-28
57
Lindner
J
P-21
96
Meyer
A
P-80
155
Lipkin
AV
P-19
94
Meyer
A
P-57
132
Liszka
B
P-76
151
Mikhailov
AM
P-60
135
Liu
YM
O-37
66
Mikhailov
AM
P-62
137
Liu
YM
P-65
140
Miki
K
P-46
121
Lo Leggio
L
P-1
76
Milathianaki
D
O-28
57
Lofstad
M
P-55
130
Misra
M
P-59
134
Logan
DT
P-64
139
Miyatake
H
P-56
131
Lu
HM
O-37
66
Monkenbusch
M
O-34
63
Lu
Q
P-50
125
Monteiro
DCF
P-72
147
Lücke
H
O-4
33
Montelione
GT
O-11
40
Luft
JR
O-11
40
Morais
M
P-10
85
Macedo
AL
P-10
85
Moreno
A
O-36
65
Machelett
MM
P-23
98
Moreno
A
P-24
99
MacPhee
C
O-22
51
Mori
Y
O-35
64
Maenaka
K
P-33
108
Morikawa
Y
P-40
115
Maertens
B
P-51
126
Moroz
OV
P-45
120
Maes
D
O-17
46
Morris
M
P-66
141
Maezawa
S
P-56
131
Müller
A
P-11
86
Maher
AH
P-1
76
Murakami
S
O-35
64
Makhov
A
O-27
56
Murata
M
P-33
108
Mäki
S
P-58
133
Nagata
Y
P-22
97
Malia
TM
P-77
152
Nagy
HM
P-13
88
Malmerberg
E
O-28
57
Nair
RR
P-8
83
Mangani
S
P-25
100
Nakai
H
P-1
76
Martin
AV
O-28
57
Nakamura
A
O-33
62
Martínez-Rodríguez S
P-48
123
Nakamura
KT
P-40
115
Martins
PM
P-5
80
Nederlof
I
O-15
44
Maruyama
M
O-35
64
Neutze
R
O-28
57
Masaki
M
P-40
115
Newman
J
O-39
68
Maschlej
M
P-43
118
Newstead
O-10
39
Matsugaki
N
P-39
114
Ng
S
J
P-66
141
Matsumoto
K
P-56
131
Nielsen
MM
O-28
57
Matsumura
H
O-35
64
Niimura
N
O-33
62
167
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no.
page
name
no.
page
Niiyama
Nikolaeva
M
AY
P-33
P-20
108
95
Plückthun
Pogoryelov
A
D
P-69
P-7
144
82
Nissen
P
O-3
32
Polidori
A
O-5
34
Nogly
P
P-71
146
Popov
VO
P-15
90
Nonaka
T
P-40
115
Popov
VO
P-19
94
Nounesis
G
P-74
149
Popov
VO
P-20
95
Oberer
M
P-13
88
Potenza
M
O-17
46
Oberer
M
P-17
92
Poulsen
JCN
P-1
76
Obmolova
G
P-77
152
Pozzi
C
P-25
100
Ogasawara
W
P-40
115
Prokofev
II
P-62
137
Ohta
K
P-40
115
Prokop
Z
P-22
97
Okada
H
P-40
115
Prudnikova
T
P-22
97
O'Keefe
S
P-42
117
Pusey
ML
P-66
141
Oksanen
E
O-32
61
Rada
H
P-13
88
Oksanen
E
P-54
129
Radajewski
D
O-20
49
Olieric
V
O-21
50
Rakitina
TV
P-19
94
Oliveira
BL
P-10
85
Raynal
S
O-5
34
Opthalage
A
O-31
60
Rengachari
S
P-17
92
Ørum
S
P-1
76
Revtovich
SV
P-20
95
Osipov
EM
P-15
90
Reynolds
S
O-27
56
Öster
L
O-7
36
Rezacova
P
P-22
97
Ostermann
A
O-34
63
Rezacova
P
P-26
101
Otrelo-Cardoso
AR
P-8
83
Rheinberger
J
P-71
146
Otten
H
P-64
139
Rho
HS
P-76
151
Otto
C
P-76
151
Ricard
F
P-28
103
Owens
R
P-13
88
Richter
D
O-34
63
Panneels
V
P-71
146
Rivas
MG
P-8
83
Passon
DM
O-23
52
Robles-Águila
MJ
P-24
99
Patel
V
P-72
147
Rocha
F
P-29
104
Pawate
AS
O-30
59
Rocha
F
P-5
80
Pearson
AR
P-72
147
Rodriguez Ruiz
I
P-48
123
Peat
TS
O-39
68
Røhr
ÅK
P-55
130
Perbandt
M
P-57
132
Romão
MJ
P-10
85
Perera
S
P-4
79
Roppongi
S
P-40
115
Perez
KS
P-24
99
Ros
A
O-18
47
Pernot
P
O-20
49
Ros
A
O-43
72
Perry
SL
O-30
59
Round
A
O-20
49
Petry
W
O-34
63
Roy-Chowdhury
S
P-4
79
Pham
NV
O-20
49
Rozov
A
P-16
91
Plentnev
S
P-6
81
Ruby
AM
O-11
40
168
name
no.
page
name
no.
page
Sakamoto
Samejima
Y
M
P-40
O-33
115
62
Sigfridsson
Silva-González
K
R
O-7
P-24
36
99
Sandmark
J
O-7
36
J
P-14
89
Santos-Silva
T
P-10
85
Singh
AK
P-68
143
Santos-Silva
T
P-8
83
Šimůnek
Singh
RP
P-68
143
Sanvito
T
O-17
46
Singh
TP
P-68
143
Saridakis
E
P-74
149
M
P-68
143
Sato
Y
P-22
97
Sinha
Sjöhamn
J
O-28
57
Sazaki
G
O-35
64
Sazanov
L
O-1
30
Scacioc
A
P-9
84
Schäfer
A
P-59
134
Schartner
J
P-11
Scherer-Becker
D
Schertler
G
Scheufler
Schieferstein
Skálová
T
P-14
89
Skálová
T
P-49
124
Skiniotis
G
O-14
43
Skråmo
S
P-55
130
86
Sleutel
M
O-17
46
P-43
118
Snell
EH
O-11
40
P-71
146
Sniady
M
P-23
98
C
P-43
118
Soltis
M
O-27
56
J
O-30
59
Sommerkamp
JAJ
P-23
98
Schindelin
H
P-59
134
Spence
J
P-6
81
Schiro
G
P-31
106
Spence
JCH
O-28
57
Schittmayer-Schantl M
P-17
92
V
O-30
59
Schlichting
I
O-28
57
Srajer
Srinivasan
VJ
O-42
71
Schmidt
B
O-39
68
Stahlberg
H
O-16
45
Schrader
TE
O-34
63
Standfuss
J
P-71
146
Schubert
R
P-57
132
Stavros
P
P-74
149
Schubert
R
P-80
155
Stellato
F
O-23
52
Schulze-Briese
C
O-21
50
Stellato
F
O-28
57
Sedzik
J
O-19
48
Sternweis
PC
P-36
111
Seetharaman
J
O-11
40
Stevenson
H
O-27
56
Seibert
MM
O-28
57
Stiens
J
O-17
46
Seine
T
O-23
52
Stojanoff
V
P-24
99
Serrano-De la Rosa LE
P-24
99
Sewell
BT
P-2
77
Sewell
BT
P-38
113
Stránský
Shah
UV
P-28
Shang
P
Sharma
S
Shiroma
J
P-49
124
Sträter
N
P-41
116
Stura
EA
P-78
153
103
Sugiyama
S
O-35
64
O-37
66
Sugiyama
S
P-33
108
P-68
143
Sun
LH
O-37
66
R
P-56
131
Sutay Kocabas
D
P-61
136
Shleev
SV
P-15
90
Suveyzdis
Y
P-11
86
Shoeman
RL
O-28
57
Suzuki
Y
P-40
115
Shuvalov
VA
P-52
127
Svensson
B
P-1
76
169
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no.
page
name
no.
page
Sweet
Syberg
RW
F
P-77
P-11
152
86
Vernede
Vettraino
X
C
P-54
P-75
129
150
Tae-Sung
Y
P-27
102
Viborg
A
P-1
76
Tae-Yeon
K
P-27
102
Viertlmayr
R
P-13
88
Takaba
K
P-46
121
Vijayaraghavan
B
P-53
128
Takahashi
S
O-29
58
Vlieg
E
O-18
47
Takahashi
Y
O-35
64
Vorontsova
MA
O-17
46
Takano
K
O-35
64
Wang
D
P-6
81
Takeda
K
P-46
121
Wang
D
O-28
57
Tanaka
I
O-33
62
Wang
TH
P-34
109
Tanaka
H
P-40
115
Warner
DF
P-38
113
Tanaka
N
P-40
115
Watts
A
O-13
42
Tang
L
O-37
66
Webb
ME
P-72
147
Tao
HR
P-63
138
Weber
BW
P-2
77
Tate
CG
O-44
73
Weierstall
U
P-6
81
Tateoka
C
P-40
115
Weierstall
U
O-28
57
Teixeira
JAJ
P-29
104
Weik
M
P-31
106
Teplyakov
A
P-77
152
Weinert
S
P-41
116
Tepshi
L
P-78
153
Welin
M
P-64
139
Terstappen
L
P-76
151
Westenhoff
S
O-28
57
Teychené
S
O-20
49
Westhof
E
P-16
91
Thaw
P
P-44
119
White
TA
O-23
52
Thunnissen
MM
P-64
139
White
TA
O-28
57
Tikhonova
TV
P-15
90
Wickstrand
C
O-28
57
Tiščenko
G
P-14
89
Wierman
JL
O-31
60
AB
P-55
130
Williams
SJ
O-41
70
Törnroth-Horsefield S
O-18
47
Williams
DR
P-28
103
Tsai
CJ
P-71
146
Williams
GJ
O-28
57
Tur
E
P-61
136
Williams
PA
O-8
37
Turano
P
P-25
100
Wilmanns
M
O-23
52
Umena
Y
P-12
87
Wilson
J
P-42
117
van der Klei
I
O-23
52
Wilson
KS
P-45
120
van Driessche
AES
O-17
46
Witkamp
G-J
P-76
151
van Enckevort
W
O-18
47
Woodhouse
J
P-31
106
Vanyushkina
AA
P-19
94
Wrenger
C
P-21
96
Vasilieva
LG
P-52
127
Wu
YF
P-69
144
Vaughn
M
P-4
79
Wu
W
P-71
146
Ve
T
O-41
70
Xie
X
P-30
105
Vekilov
PG
O-17
46
Yamada
T
O-33
62
Vera
L
P-78
153
Yamada
Y
P-39
114
Tomter
170
name
no.
page
Yang
Yano
YS
T
P-76
P-56
151
131
Yin
DC
O-37
66
Yin
DC
P-63
138
Yin
DC
P-70
145
Yin
DC
P-65
140
Yoshida
K
P-33
108
Yoshikawa
H
O-35
64
Yoshimura
M
O-35
64
Yuan
Q
P-66
141
Yusupov
M
P-16
91
Yusupova
G
P-16
91
Zahr
D
P-35
110
Zatsepin
N
P-4
79
Zatsepin
NA
O-28
57
Zhang
Y
O-17
46
Zhang
CY
O-37
66
Zhang
M
P-50
125
Zhang
CY
P-70
145
171
Did you know that all over Europe Hamburg is the city with the highest number of bridges, there are
around 2500 bridges – significantly more than in Venice, Amsterdam or London. Moreover, there is
actually a second city called Hamburg in the USA close to the Niagara Falls in the state New York.
172
The organisation team of ICCBM15 asks for your understanding, if there are minor
changes on short notice that could not have been taken into account for this conference
book. This may slightly affect e.g. the exhibition, timetable or scientific content of the
conference. We also ask you to pay attention to the local board that will keep you
updated during the conference.
We wish you a pleasant and fruitful event.
Hamburg, 2014
A FEW HELPFUL GERMAN PHRASES
Hello
Hallo
Goodbye/see you
Auf Wiedersehen
Thank you
Dankeschön
Yes/No
Ja/Nein
How are you?
Wie geht es dir? (informal),
Wie geht es Ihnen? (formal)
I am lost.
Ich habe mich verirrt.
Where is…?
Wo ist…?
How much does it cost?
Wieviel kostet es?
Long time no see.
Lange nicht gesehen. (informal)
I don’t understand.
Ich verstehe nicht.
I need help.
Ich brauche Hilfe.
I am just kidding.
Ich mache nur Spaß.
Reservation
Reservierung
Menu
Speisekarte
Ticket machine
Fahrkartenautomat
Excuse me!
Entschuldigen Sie!
173

Folie 1

Transcription

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