PDF of the 2010 Nematode Lab Manual

NEMATODE MODULE
MBL EMBRYOLOGY 2009
Faculty:
David Fitch1
Paul S. Maddox2
Amy S. Maddox2
Joel Rothman3
David Sherwood4
Teaching Assistants:
Elliott Hagedorn4
David Q. Matus4
1
Department of Biology, New York University, Main Building, Room 1009, 100 Washington Square East
Greenwich Village, New York, NY 10003
2
Institute for Research in Immunology and Cancer, Department of Pathology and Cell Biology, P.O. Box
6128, Station Centre-Ville, Montreal, QC H3C 3J7, Canada
3
Department of Molecular, Cellular and Developmental Biology, University of California, Santa Barbara, CA
93106
4
Department of Biology, Box 90338, Duke University, Durham, NC 27708
I.
Organization of the Lab
II.
Introduction to C. elegans
A. Overview
B. Establishment of embryonic polarity in C. elegans
C. Vulval induction in the larva
D. Male mating behavior
E. Ovulation
F. Basement Membranes
G. Comparative development: Natural variation in species related to C. elegans
III.
Techniques Used in this Laboratory
A. Basic worm handling
1. Picking worms
2. Mounting embryos/larvae for Nomarski and fluorescence microscopy
3. Obtaining early embryos
B. Laser cell ablation and fusion
1. Cell ablations
2. Cell fusions
C. Green fluorescent protein (GFP) markers
D. RNAi by ingestion of E. coli expressing dsRNA
IV. Experiments
A. RNAi screen
1. RNAi screen for new genes that regulate uterine-vulval development
B. Ablations/Fusions
1. Ablations to examine cell lineage, cell-signaling, inductions, etc.
2. Vulval & gonad development in C. elegans & different species
C. Male mating behavior
1. View male mating
2. View Sensory neurons by labeling with DiI
3. View male mating in pkd-2 mutant
4. View male mating in different species
D. Wild nematodes
E. Live cell imaging
1. Transmitted light imaging
2. GFP-fusion proteins during the first cell division
3. View tissue specific marker strains
4. Observing male tale development in different species
5. Live analysis of sperm behavior
6. Ovulation time-lapse
7. Using photoconversion to track basement membrane dynamics
8. Vulval development in other species (an evo-devo project)
V. Lab Schedule
VI.
Appendices
A. Strain list
1. Mutant strains
2. Other nematodes
3. Strains expressing tissue-specific GFP markers
B. Founder cell lineage and cell fate markers
C. Sensory organs of the male tail
D. Gamete maturation and oöcyte ovulation
VII. References (electronic copy only)
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I.
Organization of the Lab
We have six formal lab periods beginning on Thursday afternoon. During the orientation period,
we will review techniques that you will need to master in order to perform your experiments
(manipulating worms; mounting embryos, etc.). You will be introduced to a number of major
experimental procedures commonly used in C. elegans embryology laboratories: RNA
interference (RNAi), laser cell ablation/fusion, and expression analysis by GFP. We have
available much more than can readily be done by one student and a number of new experiments
for which the outcome is unknown. Keeping this in mind as you read through this manual,
choose experiments you are the most interested in pursuing. The three David’s, Joel, Elliott,
Amy, and Paul will be happy to assist you with experiments in order for you to become supreme
handlers of “the worm”!
II. Introduction to C. elegans
A. Overview
C. elegans, a free-living soil nematode, was chosen by Sydney Brenner in 1963 as an
experimental organism because it is transparent, easy to propagate and is amenable to genetic
manipulation. The entire cell lineage of C. elegans, from the zygote to the adult, has been
determined. Exactly 1090 somatic nuclei are produced during its development to adulthood; 131
cells undergo programmed cell death; adults contain 959 somatic nuclei. Its transparency and
reproducible anatomy make it possible to identify each cell at all stages of development; the
lineage provides us with the knowledge of the ancestry and ultimate fate of every cell. These
features, coupled with its amenability to forward genetic, reverse genetic and molecular studies,
have allowed developmental, physiological and behavioral events to be identified and
characterized at an unprecedented level. The scientific contributions of this small worm are
highlighted by the Nobel Prize for Physiology or Medicine award in 2002 to Sydney Brenner,
Robert Horvitz and John Sulston for their characterization of organ development and
programmed cell death in C. elegans, the 2005 award to Andrew Fire and Craig Mello for their
characterization of RNA-mediated interference, or protein depletion and the 2008 Nobel Prize in
Chemistry to Martin Chalfie for his use of the green fluorescence protein (GFP) as an
experimental reagent in C. elegans.
C. elegans exists as two sexes: self-fertilizing hermaphrodites, which produce both
sperm and oöcytes, and males. Self-fertilizing hermaphrodites make it possible to propagate
large numbers of animals without the necessity of mating with males. Thus, many worms can be
generated by placing a single hermaphrodite on a petri dish with sufficient food (worms are
grown on a lawn of E. coli). Genetic crosses are performed by mating males with
hermaphrodites; in such a mating, the sperm from the males are preferentially used and the
hermaphrodite serves the role of a female.
In this lab module, you will have the opportunity to observe the development of wildtype worms and mutants defective in different aspects of development and behavior. You will
also carry out laser ablation (and/or fusion) of identified cells, analysis of differentiated cell
types by green fluorescent protein (GFP) markers, determination of cell lineages, and reverse
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genetics by RNAi. One of the main advantages of C. elegans for genetic studies is its short
generation time (3 days). Although the constraints of a short module will not allow sufficient
time to perform genetic experiments, you will have the chance to set up a mating and observe
mating behavior.
B. Establishment of embryonic polarity in C. elegans
The first cell division in Caenorhabditis elegans is asymmetric. Fertilization not only instigates
the completion of maternal meiosis, but also supplies the cell with a single pair of centrioles,
which separate to form two spindle poles shortly after sperm entry. These two events precede
the dynamic movement of cortical cytoplasm from the posterior towards the anterior, and central
cytoplasm moving from the anterior end towards the sperm pronucleus in the posterior. These
cortical and central cytoplasmic flows occur for about 7 minutes, roughly at the same time that
cell fate determinants, such as the posteriorly positioned germ line P granules, become localized
to the posterior. Disruption of the actin cytoskeleton interrupts polarized cytoplasmic flow and
the contents of the first cell are not asymmetrically localized. A group of genes known as the
partitioning defective (par) genes are required for asymmetric division of the zygote.
C. Vulval induction in the larva
Vulval induction in C. elegans has provided an important model for examining cell fate
specification. The entire vulva is derived from a single 1o and two 2o-fated vulval precursor cells
(VPCs), which lie along the ventral epidermis. The 1o and 2o VPCs give rise to lineages of eight
and seven cells, respectively, which have distinctive division and gene expression patterns.
Specification of these VPCs to a vulval fate is dependent on LIN-3, a protein similar to
mammalian epidermal growth factor. LIN-3 is produced by the anchor cell (AC), a specialized
cell in the developing gonad, during the late L2 to early L3 larval stage. LIN-3 activates a
receptor tyrosine kinase pathway in the nearest VPC, P6.p, specifying it to adopt a 1o fate. One
aspect of the 1o fate specification is the upregulation of three distinct ligands for the C. elegans
Notch receptor LIN-12. This upregulation directs the neighboring VPCs, P5.p and P7.p, to adopt
the 2 o-fate. Without the inductive LIN-3 signal from the AC, no vulval induction occurs.
Instead all VPCs divide once and then adopt a non-vulval 3o fate and contribute to the external
epithelium that covers the animal. The resulting vulva-less phenotype can be used as a genetic
tool: Conditional disruption of expression of genes required for vulval specification blocks
vulval formation, and hermaphrodites lacking a vulva cannot lay embryos. Thus, any embryos
that are produced hatch inside the mother and eat the body, causing an easily-scored “bagging”
phenotype.
D. Male Mating Behavior
C. elegans is an outstanding model system for behavioral analysis. The simple wellcharacterized nervous system, complete genome sequence, short generation time and amenability
to genetic analysis facilitate detailed study of behavior at the organismal, cellular and molecular
levels.
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Male mating is a complex behavior in C. elegans, which can be divided into the following simple
steps:
1) Response and backing: When the posterior part of a male comes in contact with a
hermaphrodite, the male responds by placing the ventral surface of his tail against her body and
moves backwards, searching for the vulva.
2) Turning behavior: If the male reaches the end of the hermaphrodite (either head or tail), he
will turn and continue to back along the other side.
3) Vulval location: In most cases, the male will stop backing when he scans over the vulva.
4) Sperm insertion: The male inserts his spicules into the hermaphrodite.
5) Sperm transfer: The male then transfers his sperm into the hermaphrodite.
response
turning
vulval location
spicule insertion
Figure 2.1 Male mating behaviour
E. Ovulation
During oöcyte maturation in C. elegans, the oöcytes align along the proximal-distal axis of the
gonadal sheath and mature in an assembly-line manner as they proceed proximally toward the
spermatheca (see appendix). Ovulation is the exit of the most proximal oöcyte from the gonad
arm into the spermatheca. This process requires the contraction of the six proximal myoepithelial
sheath cells and the dilation of the distal spermatheca. The sheath contractions pull the dilating
distal spermatheca over the oöcyte, placing the oöcyte into the lumen of the spermatheca where
fertilization occurs. The fertilized egg then moves from the spermathecal lumen to the uterine
lumen through the spermathecal-uterine valve. Fertilization triggers the exocytic events that
promote eggshell formation, and after this point, the fertilized egg takes on its characteristic
oblong “egg shape.” Before this, the large oocyte is remarkably deformable, bounded only by a
plasma membrane, and squeezes through the tiny valves.
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F. Basement Membranes
One of the defining structures present in metazoans is basement membrane. This specialized
form of extracellular matrix is thin (60-120 nm), dense and highly cross-linked. Its mechanical
strength provides the structural underpinning for all epithelia, endothelia and many mesenchymal
cells. Basement membranes are also important for organizing tissues into distinct compartments,
tissue repair and in guiding migrating cells during development. About 50 proteins are known to
make up the basement membrane. The main components of basement membranes include type
IV collagen, laminin and nidogen. The major basement membrane components in vertebrates
are all present in C. elegans. Furthermore, in C. elegans these proteins are encoded by much
smaller gene families. For example, laminins are heterotrimeric proteins composed of a
α, β, and γ chain. C. elegans have two α chains , and a single β and γ chain, forming just two
laminin isoforms compared with 15 in vertebrates. Thus, C. elegans is particularly amenable to
genetic analysis of basement membrane formation and function.
G. Comparative development: variation among species related to C. elegans
An important question concerning a "model system" is to what degree are the characters
(features) actually representative of other species. Beyond this, however, variation can be used
to help understand how characters have evolved, how evolving systems adapt to environmental
situations, and what elements of developmental systems are constrained from changing or can
change without deleterious effect to the system. Comparative studies are best done in the context
of a phylogeny, which allows one to infer the direction and number of evolutionary changes. C.
elegans is one of many related free-living species ("rhabditids") that differ to varying degrees in
terms of many characters. For example, some species have a vulva that is positioned in the
midbody; others have posterior vulvae. Many of those with a midbody vulva (like C. elegans)
require an induction signal from the gonad to produce the vulva from ventral Pn.p cells; many
with a posterior vulva are induction-independent. Early cellular events during embryogenesis
also vary substantially. The male tail varies in terms of positions of the rays, the shapes of the
spicules and fan, and the shape of the tail tip, all characters that have been classically important
for taxonomy and phylogenetics. Thus, rhabditids can be used as a powerful system to study
both how developmental mechanisms underlying a conserved feature can change as well as how
development changes to make different forms.
III.
Techniques Used in this Laboratory
A. Basic worm handling
1. Picking up worms. Flame the platinum wire of your pick until it glows red and let it cool
briefly in the air. Put a glob of bacteria on the end of the wire (you may find it helpful to take it
from a worm-free plate, e.g., the one to which you will be transferring worms). Touch the glob
to a worm, using the sticky bacteria to pick up the worm. (Don't "scoop" the worm; just touch the
flattened underside of the pick to it.) Then set the worm down on a new plate in the thick part of
the E. coli lawn. You should try not to gouge either plate as this will encourage the worms to
burrow into the agar, where they become inaccessible. Start by transferring one worm at a time;
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eventually it is possible to carry 10 or more worms on one pick-full. Be patient; it takes practice
to get used to transferring worms. Sometimes worms don't like to come off a pick easily; in this
case, soften the glue glob by moving the loaded pick back and forth gently in the E. coli of the
plate to which you are transferring.
TO DO:
Learn how to identify (1) eggs, (2) L1 larvae, (3) L4 larvae, and (4) adults on your
N2 plates. Practice transferring worms from plate to plate with a pick.
Eggs are oval and easy to identify. Adults are the largest worms on the plates and carry
eggs visible in the middle of the ventral side. (Worms crawl on their side, not their ventral
surface.) L1 larvae are the smallest things crawling on your plates. They are sufficiently small
to fit inside an eggshell, which they recently did. At their youngest, they have no dark coloring,
unlike other larvae and adults. L4 hermaphrodites are best identified by their invaginated
developing vulvae that look like pale half moons on the ventral side in the middle of the worm.
There is a black dot in the half moons (see Figure 2.2 below) in L4 larvae.
Figure 2.2 L4 hermaphrodite
On the video display connected to the Nomarski scope, we will look at oogenesis and
early embryogenesis in an intact adult hermaphrodite. We will point out some cells and
structures you will need to identify for yourselves for laser ablations, and we will cover such
basics as identifying left/right and dorsal/ventral. Feel free to go back and forth between
anatomy and lineage charts.
2. Mounting live embryos or larvae for Nomarski (DIC) microscopy. You will have the
chance to view the development of a newly fertilized zygote and perform laser ablations of cells
using the Nomarski scopes. The following outlines the general steps. The techniques are
discussed in more detail below.
To prepare a Nomarski slide you need to:
1. identify the desired stage of embryo, zygote, or worm
2. transfer it to an agar pad on a slide
3. add a cover slip
4. seal with Vaseline or Valap (1:1:1 vaseline, lanolin and paraffin) if you are doing timelapse imaging.
TO DO in advance:
1. Ensure that tubes of agar (and Valap) are thoroughly melted.
2. Prepare a capillary tube for your mouth pipette for liquid transfer of eggs.
Pull a capillary over a flame so that the end is about the diameter of an adult worm (we
have also prepared pre-pulled capillaries for you). You want it to be big enough to easily suck
up eggs and larvae, but small enough that it can form a seal with a drop of water. Once you
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obtain a capillary to your liking, it can be used for all of your experiments. Attach it to a mouth
pipette.
2 layers of tape
molten agar
agar pad
4th slide
guide slides
Figure 2. Set-up for preparing agar pads
Prepare an agar pad by placing a generous drop of molten 5% Noble agar on a plain
microscope slide. This slide should be placed between two guide slides whose surfaces have
been raised slightly with two layers of tape so that the agar pad has the proper thickness (Fig. 2).
Work quickly so the agar does not solidify. Perpendicular to these slides, place a fourth slide
over the agar drop so that a pad is formed. Do this by placing one side of the fourth slide at the
edge of the molten agar, avoiding trapping air bubbles in the agar. Press the top slide down
against the supporting slides to flatten the agar. Wait a couple of minutes for the agar to solidify.
Once the desired specimens are ready to mount, lift the two slides contacting the agar
away from the guide slides. Carefully tease the two apart by a back and forth horizontal motion
that will leave the agar pad on one slide only. Do not rip the pad as you tease the slides apart.
Try to work steadily now so that the agar pad does not dry. (You can pipette some H2O around
the edges of your pad to keep it moist).
You want to transfer your embryos in a very small volume of liquid. Using the mouth
pipette, take up M9 or egg medium into the capillary (no more than 1-2 microliters or your
embryos will float away!!). Trim the agar pad to approx. 8 mm x 8 mm with a razor blade. Suck
up your eggs up into the capillary and gently blow them in a small volume of liquid onto a corner
of the trimmed agar pad. Using an eyelash mounted to a toothpick, cluster the embryos by
sliding them to the center of the pad. WORK QUICKLY AND KEEP THE EMBRYOS
MOIST. IF THE WORMS DRY OUT THEY WILL DIE. Using your fancy forceps, gently
cover the embryos with a 22 x 22 mm coverslip and seal with melted Vaseline held in the heat
block (back bench in lab).
To transfer larvae or adult worms to a slide you first add a drop (2.5µl) of M9 on top of
your agar pad. Collect worms as outlined above with your platinum wire, and then gently move
the glob of bacteria containing your worms side-to-side in the drop of M9. This should free most
of the worms from the bacteria. Using your forceps or fingers, gently cover the embryos with a
22 x 22 mm coverslip and seal with melted Vaseline.
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3. Obtaining early embryos. Transfer about 20 adult hermaphrodite worms to a drop of about
400 microliters of M9 in a depression well. (We will not be using it here, but often researchers
use a special egg medium if embryos younger than the 2-cell stage are desired). Take two 25
gauge needles and, using them like scissors, cut the hermaphrodite in half near the vulva. This
will release eggs into the medium. Pulling the cut worms in and out of your mouth pipette will
also help to dislodge embryos. Collect or cluster the embryos of the desired stage either with
your mouth pipette or with an eyelash. (See us for needles if you are having trouble pulling the
right size). Using your mouth pipette, transfer the desired embryos to the agarose pad and cover
with a coverslip. Seal with Vaseline to prevent dehydration.
B. Laser Cell Ablation and Fusion
Laser microsurgery provides a precise and rapid method for eliminating or fusing cells in
the animal. A pulsed dye laser is arranged as an epi-illuminator, its beam being directed
downward through the objective by way of a semi-silvered mirror. Accurate targeting is
achieved by pre-alignment of the focal point of the laser with a reference reticule in the eyepiece.
We will divide the class into three groups for demonstration; after this instruction you can
sign up for individual time on the laser. If your scheduled or desired time conflicts with
someone else’s, consider that while one person is ablating, another person can be setting up their
next slide. Normal, rigorous technique would require that you regularly monitor your animals
after ablation. This is not practical given our time and equipment constraints. Also, you should
think about the caveats of laser ablation experiments. How do you know when to ablate a cell?
What is the significance of a negative result? What happens to the cytoplasm and cell membrane
after you ablate a cell and how might this affect (or not affect) communication between cells?
1. Cell Ablations. Careful control of the laser pulse power is necessary when performing cell
ablation. Too little power has no effect, and too much power causes excessive damage, often
bursting the embryo. Generally, the best strategy is to deliver several sub-threshold pulses
within a target area. For most ablations, the beam is directed at the nucleus of the cell to be
killed. Usually, debris can be seen to appear in the nucleus after several successful hits. The
beam is directed at various points throughout the nucleus by moving the stage in the x, y, and z
axes. Often, cells that are markedly damaged after ablation will recover and go on to divide; it is
therefore advisable to observe the ablated cell for some time to ensure that it dies.
2. Cell fusions. Cell fusions can be performed to combine the cytoplasmic contents of
neighboring cells. Cell fusions are performed by directing laser pulses to the apposing
membranes of the cells to be fused. Only a few hits are required to completely fuse two cells;
this procedure requires more practice than cell ablation. In most cases a tetrapolar cleavage
occurs during the subsequent cell cycle, generating four mononucleate cells from the fused
dikaryon, all containing cytoplasm from both of the fused cells.
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C. Green Fluorescent Protein (GFP) Markers
The green fluorescent protein (GFP) from the jellyfish Aequorea victoria has proven to be an
extremely useful reporter protein in gene expression constructs, since it is visible in living
animals. So useful, in fact, the 2008 Nobel Prize in Chemistry was awarded to Osamu
Shimomura, Martin Chalfie and Roger Y. Tsien for discovering and developing GFP as a
experimental reagent. GFP fluoresces green when illuminated directly with blue light.
Therefore, gene expression can be assayed directly in living worms. This provides us with the
ability to follow dynamic events during development and eliminates the need for fixation and
antibody or in situ staining procedures. The GFP markers can be used in the laser ablation
experiments as well as for visualizing tissue-specific gene expression patterns. To view GFP,
mount embryos as for Nomarski microscopy and observe under the FITC or a narrow-band
GFP(S65C)-specific fluorescence filter. The strains that contain GFP reporters are listed in
Appendix A.
D. RNAi by Ingestion of E. coli expressing dsRNA
The technique of RNAi (RNA-mediated interference) allows an assessment of a loss-of-function
phenotype from a gene of interest virtually overnight; it has revolutionized studies of gene
function in C. elegans and more recently in other organisms as well. After delivery of doublestranded RNA (dsRNA) to nematodes, phenotypes can be observed in both the treated animals
and in the progeny. Remarkably, the RNAi effect with many genes is potent even when worms
are simply soaked in a solution of dsRNA, or grown on a strain of E. coli expressing sense and
antisense RNAs (which anneal in vivo to make dsRNA). Such ‘feeding plates’ contain a lawn of
concentrated E. coli HT115 harboring a plasmid in which convergent T7 promoters flank an
inserted cDNA fragment. Transcription of both strands occurs through IPTG induction of a
lacO::T7 RNA polymerase chimera.
IV. Experiments
With the exception of the RNAi experiment, you will have your choice of which experiments
from this list to choose from. There are far too many to do them all. We want you to pick the
ones that sound the most interesting to you. Some experiments are more difficult than others
and you might discuss them with one of us before attempting them. To make your lab time most
productive, you should design your experiments prior to the lab period. This is especially true
for the ablations and free periods. Consider what you may need to prepare in advance. For
example, if you want to examine adult hermaphrodites after RNAi, pick the appropriate staged
worms for the treatment.
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A. RNAi—You will work in pairs on the RNAi experiments
Experiment: RNAi screen for Genes that Regulate Uterine-vulval Development:
Uterine-vulval development in C. elegans provides a powerful model to study animal
organogenesis, as it involves a limited and stereotyped number of cells, occurs in less
than 24 hours, and can be approached with a unique experimental toolkit that includes
live-cell imaging, forward and reverse genetics, and a host of molecular methods. Classic
studies of uterine-vulval development have had a profound influence on our current
understanding of conserved signaling pathways like Wnt, EGF-Receptor Tyrosine
Kinase, and Notch.
Formation of the vulva (which provides a connection between the uterine tissue and
the outside world) involves a series of cell migrations, cell shape changes, cell-cell
attachments, and cell-cell fusions. These cellular processes are spatially and temporally
controlled by a precise coordination of intercellular signaling, signal transduction, and
transcriptional regulation. During the first larval stage, 12 ventral hypodermal cells are
born (P1-12).p, six of which (P(3-8).p) comprise what is called the vulval competence
group (they have the capacity to become vulval cells). A single cell in the somatic gonad,
called the anchor cell (AC), then secretes an EGF-like ligand that induces the nearest
three vulval cells to adopt a vulval cell fate. Via a second lateral Notch signal between
these vulval cells, the cell closest to the AC (P6.p in wild-type; directly beneath the AC)
will adopt a primary fate and divide to generate 8 cells. The two cells next to P6.p (P5.p
and P7.p) will adopt a secondary fate, dividing to form 7 cells each (Fig. 3A). P(3-4).p
and P8.p adopt a tertiary fate where they divide once and fuse with the hypodermis. The
resulting 22 vulval cells will eventually give rise to the vulval tissue of the adult
hermaphrodite after a series of cell fusions. Shortly after the AC induces the vulval fate,
it plays a second role in uterine-vulval development by invading through the BMs that
separate the uterine and vulval tissues, initiating the connection between these tissues
(Fig. 3A).
Specific defects in uterine-vulval development can result in egg-laying defective (Egl)
and protruded vulval (Pvl) phenotypes, both of which are visible at the plate-level and
can be scored using a standard stereomicroscope. Thus, one can screen for genes
involved in uterine-vulval development by first scoring for Egl or Pvl phenotypes and
then subsequently using Nomarski optics and fluorescence markers to further investigate
the underlying defect. Among the defects that give rise to Egls or Pvls are improper
vulval cell fate specification, a failure in uterine-vulval connection, and disruption of the
cell-cell fusions that take place.
Genome-wide RNAi screens in C. elegans have identified hundreds of genes that give
Pvl or Egl phenotypes when targeted by RNAi. We have selected 12 of these genes that
have not been previously characterized in uterine-vulval development. Your job will be
to examine available fluorescence marker strains (transgenic worms) that have been
treated with RNAi targeting one of these genes and see if you can determine the
underlying defect in uterine-vulval development. The defect underlying the Pvl or Egl
phenotype after knockdown of these genes is unknown. You may find specific defects in
uterine-vulval cell fate specification, uterine-vulval attachment, cell-cell fusions, etc.
Alternatively, the defect may be in tissues for which markers will not be available. Thus,
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you may be limited in your ability to identify the defect using the marker strains you are
given. These marker strains will include a vulval cell fate marker (egl-17>GFP), a
uterine cell fate marker (cog-2>GFP), and an AC-specific F-actin marker
(mCherry::moeABD) with a basement membrane marker (laminin::GFP) (Fig. 3B-D, see
appendix for complete description). You will pair up with your neighbor and be given a
6-well plate that already has the worms (the afore mentioned fluorescence marker strains)
plated on bacteria that express RNAi targeting one of the 12 genes. These plates will also
contain wells with bacteria expressing the empty RNAi vector to serve as a negative
control for each strain.
Figure 3. Visualizing uterine-vulval development.
(A) Schematic diagram shows several stages of uterine-vulval development. Left panel represents the P6.p two-cell stage of the mid
L3 stage, after P6.p has divided once. Middle panels depict the P6.p four- and eight cell stages, specific cell fates are shown for each
tissue. The right panel shows the mid L4 stage when the AC fuses with the neighboring uterine cells. (B) Image shows double
transgenic strain expressing the F-actin marker (mCherry::moeABD) specifically in the anchor cell (AC) and laminin::GFP in the
basement membrane (BM) at the P6.p four-cell stage. By this time, the AC has created a gap in the underlying BM (see arrows,
middle panel). (C) Images show animal expressing nuclear localized cog-2>GFP in the uterine cells that neighbor the AC (shown in
magenta). (D) Image shows animal at the P6.p eight-cell stage expressing egl-17>GFP in the central vulval cells. In this strain, the
BM is labeled with laminin::mCherry. See appendix for more detailed descriptions of marker expression patterns. Images were
kindly provided by Dr. Shinji Ihara, Duke University.
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Outline:
•
•
•
•
•
•
We have a 6-well RNAi feeding plate
available for pairs of students to examine
RNAi targeting one of the 12 genes with
three different fluorescence marker
strains.
The left wells (3) of each plate contain
your controls.
These wells contain
empty vector (L4440) control RNAi.
The right wells (3) have RNAi targeting
one of the 12 genes.
Each row contains a different
GFP/mCherry strain to visualize a
different aspect of uterine-vulval
development.
Wednesday evening, your TAs plated
~100 arrested L1 stage worms in each
well of your 6 well plate. By Thursday
evening these worms should reach the
late L3 / early L4 stage, when you can
begin to determine what defects the
RNAi might be causing. Your TAs will
re-plate the same experiment for you on
Thursday evening so that you can repeat the scoring again on Friday.
You can being by looking at your L4440 empty vector controls under high magnification so
you will know what wild-type development looks like. Then we would like for you to
compare the RNAi knockdown to the L4440 negative controls for each strain.
We will generate an excel document for everyone to input their scoring data into so that it
will be available to everyone the class.
Protruded Vulval Phenotype (Pvl)
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Schedule:
(TAs)
Wednesday evening
Thursday afternoon
(Students)
Thursday
Friday
Plate L1 arrested larva on 6 well RNAi plates (round 1)
Plate L1 arrested larva on 6 well RNAi plates (round 2)
Students score RNAi plates for uterine and vulval defects (round 1)
Repeat experiment (round 2)
Exercises:
a) Look up your gene targeted by RNAi on Wormbase: Wormbase is an
integrated database that incorporates comprehensive information about C.
elegans, including all genomic data, literature, genetics, etc. We will demonstrate
how to use this database, which you can use to look up your gene, find the protein
domains/similarities, etc.
b) Scoring for uterine and vulval defects: At the beginning of the lab section we
will demonstrate what wild-type expression for each GFP/mCherry reporter strain
looks like. Following this you can look at both wild-type expression (L4440
empty vector controls) and whether the RNAi depletion affects any of the reporter
strains. This scoring should be done at high magnification (63X or 100X
objectives).
c) Repeat scoring for uterine and vulval defects: The TAs will plate a second
round of worms on fresh 6-well plates on Thursday evening so that you may
repeat the experiment Friday evening.
B. Ablations/fusions
1. Ablations to examine cell lineage, cell-signaling, inductions, etc.
a. Ablation of MS. One example of an interesting ablation is the ablation of MS. This
prevents formation of the pharynx (since the posterior pharynx is produced by MS and the
anterior pharynx is induced in AB by MS) and eliminates left/right differences in the AB
lineage, resulting in dramatic cell lineage alterations. You can use the ceh-22::GFP strain
for these experiments to follow expression of ceh-22 (restricted to the pharynx).
b. Laser-induced fusions. Fuse oöcytes to make giant embryos. Fuse blastomeres to ask
which fate is dominant when two cells are made into a heterokaryon.
14
2. Vulval & Gonad Development
The anchor cell (AC) has a key role in inducing the vulva during the L2 stage, but also has
a more subtle role in vulval patterning during the early L3 stage
a. Laser ablate the anchor cell (AC) in syIs50 (cdh-3::GFP is expressed in the AC) at early
L3 stage. Is vulval morphology normal? If you save animals with perturbed or no vulval
induction what happens to the embryos trapped within the animal?
b. Laser ablate a distal tip cell at the end of one of the gonad arms during the L2 or early
L3 larval stage or in the adult stage. Recover the worm and let it grow on a plate overnight.
What happens to the gonadal arm where the distal tip cell was ablated?
c. Laser ablate the linker cell in the male. Recover the worm and let it grow on a plate
overnight. What happens to the males gonad when the linker cell is ablated?
d. Laser ablate the AC or somatic gonad precursors (Z1 and Z4) in other species: e.g.
Pelodera strongyloides (which has a midbody vulva) and Teratorhabditis palmarum
(which has a posterior vulva). Try to recover the worms and let them grow for a couple
days to adulthood. Which ones require induction from the gonad to make a vulva? How
might you distinguish between experiments showing independence from induction and
experiments in which your ablation may not have worked?
C. Male mating behavior
1. View male mating
Pick L4 stage males onto an NGM (nematode growth medium) OP50 bacterial seeded
plate and allow males to develop overnight. Pick L4 unc-31 hermaphrodites onto an
NGM seeded plate and allow to develop for 3 days (we'll pick these ahead of
time…..older, paralyzed hermaphrodites are used because males can mate and insert
spicules into these females more easily). Pick 5, three-day old, unc-31 hermaphrodites
onto an NGM seeded plate with a small spot of bacteria. Place a single male near the
hermaphrodites and see if you can identify the different steps in mating.
2. View sensory neurons by labeling with DiI
With their exposed external cilia, certain chemosensory and mechanosensory neurons
take up dye in the environment, allowing these neurons to be specifically labeled. The C.
elegans male tail has several neurons that can be labeled using DiI. You may also like to
stain males from other nematode species with DiI; you can then compare the organization
and morphology of these neurons in different species.
15
Protocol:
a. DiI Stock solution is 2 mg/ml in dimethyl formamide, stored at –20oC in a foil wrapped
tube.
b. Dilute the stock 1:200 in M9. Some dye will precipitate when you do this; don't worry
about it.
c. Put 150µl dye solution in a microtiter well, and use a worm pick to transfer some
worms into the dye. Incubate 2-3 hours at room temperature.
d. Use a mouth pipette to transfer the worms to a fresh plate, and let them crawl on a
bacterial lawn for about 3 hours to destain.
e. Put worms on pads with 0.2% tricaine; 0.02% tetramisole in M9 or 1 mM azide and
visualize by fluorescence using the appropriate filter.
3. View male mating in pkd-2 mutant
Observe mating in a pkd-2 (polycystin-2) mutant and view the expression pattern of a
pkd-2::GFP reporter construct.
pkd-2 encodes a putative channel protein and is the C. elegans homologue of the human
polycystin-2 gene, a gene mutated frequently in polycystic kidney disease.
Is the
reporter construct expressed in hermaphrodites? Is there anything curious about the
male expression pattern? In what step(s) in mating are pkd- 2 mutants defective?
4. View male mating in different species
Observe mating on plates using a dissecting microscope. What mating positions are
adopted by the different species? Are there preferences for a particular orientation (e.g.
parallel versus antiparallel, left coiling versus right coiling)? Map the mating positions
onto the phylogeny to determine which state is ancestral versus derived. What factors
may contribute to determining mating position? (Can you make any phylogenetic
correlations?)
Can you distinguish between pre- and post-zygotic reproductive barriers between
species? You could set up interspecific mating tests and determine if mating occurs, if
hybrid eggs are laid, and if viable F1s are produced. However, be certain you pick
unfertilized virgins for your experiments. (If a species is a self-fertile hermaphrodite,
mate them only after they have exhausted all their sperm.)
16
D. Wild Nematodes
Nematodes can be collected from the wild outdoors by taking an empty plate and
sprinkling a small amount of soil from outside onto a fresh agarose plate with E. coli.
After putting soil on the plate, replace the lid, and place the plate on your bench. As soon
as a few hours after putting the soil into the plate, Nematodes can be seen crawling on the
lawn of E. coli. You should transfer these worms to fresh plates. Next, you can collect
and mount embryos to compare early cell divisions. Collect and mount early embryos as
you would with C. elegans to do timelapse recording of the first cell divisions. Are the
first cell divisions asymmetric, as with C. elegans? If they are symmetric, how do you
think localization of P granules occurs? When is the germline specified? What simple
experiments could be done to determine when P granules are segregated and when the
germline is specified? The ablation and mating behavior experiments suggested above
can also be performed on these species.
Caenorhabditis species are not really "soil" nematodes, but have been found in
different kinds of habitats. Elegans-group species, such as C. elegans, C. briggsae,
C. remanei, and some as yet unnamed Caenorhabditis species have been found in
compost heaps, with reproductive adults found on freshly rotting fruit. (Maybe they
should be called "fruit nematodes"!) Other Caenorhabditis species have been found in
association with rotting cactus (C. drosophilae, C. sonorae), dens of burrowing bugs (C.
japonica) and in phoretic associations (generally as dauer larvae) with snails, pill bugs
(Armadillidium) and millipedes. If you are interested in trying to discover new
Caenorhabditis species (there should be many more out there), you could try setting bait
"traps" with fresh fruit near a compost pile or other place; after a few days, there will be
nematodes on your now rotting fruit. Check them for distinguishing characteristics of
Caenorhabditis or Elegans-group species (e.g. prominent median bulb in the pharynx, fat
ray 6 in the male tail, "hook" structure on the precloacal sensillum).
E. Live cell imaging
1. Transmitted light imaging. Many important events in the early embryo can be
observed by transmitted light imaging. The cytoplasm is largely filled with refractile yolk
granules, which bear witness to the flows that occur during the establishment of anteriorposterior polarity. The cortical cytoplasm flows towards the anterior, and the central
cytoplasm flows towards the posterior. The nuclei are clearly visible as round clearings in
the granular cytoplasm. The sperm pronucleus, initially residing in and defining the
posterior, expands from 2-4 microns compacted to 7-10 microns at maximum. This
characteristic increase in size is an indication of cell cycle progression. Nuclear envelope
breakdown is visible. The centrosomes, or microtubule organizing centers of the mitotic
spindle, are also visible as clearings in the cytoplasm, and their increase in size is also
characteristic of mitotic progression. It is possible to see yolk granules and other features
aligned along the long straight microtubules (themselves invisible) that emanate from the
centrosomes towards the cortex.
17
One main strength of using the C. elegans early embryo as a model system for cell
biology is the reproducibility of timing, placement, magnitude, etc of events. You can
perform time-lapse imaging of several embryos and measure a developmental event and
compare the timing, etc. among your specimens.
2. GFP-fusion proteins during the first cell division. Using the method described
above for dissecting embryos for live cell imaging, isolate embryos from the appropriate
transgenic strain. (For example, observation of chromosome dynamics requires TH32,
GFP:Histone H2B and GFP:gamma tubulin). We will also have fluorescent markers for
components of the cortical actomyosin cytoskeleton. Image the embryos by DIC and
fluorescence using widefield or spinning disk confocal microscopy. Record several
untreated (control) divisions prior to timelapse recording sequences from RNAi treated
worms to ensure a well-controlled interpretation. See the microscopy handbook for
details on setting up the microscopes and proper alignment. In untreated embryos, note
the timing and rates of nuclear migration, nuclear envelope break-down (NEBD), spindle
orientation, anaphase onset, cytokinesis onset and completion, as well as the relative sizes
of the resulting blastomeres.
We will have available RNAs directed against genes whose products are implicated in
cell division. You could treat L4 stage worms with soaking RNAi to deplete target
proteins from the gonad and resulting embryos. Compare cell division parameters in
control and RNA-treated embryos and devise a quantitative method to characterize the
role of the RNAi target protein in the first cell division.
3. View tissue specific marker strains. In the appendix is a list of transgenic strains
with GFP expressed in different tissues. These strains are available at the front of the lab,
and be easily mounted and viewed for fluorescence.
4. Observing male tale development. Pick early L4 males and mount them onto agar
pads with levamisole anesthetic. Watch morphogenesis occur over a period of 2-4 hours.
This can also be done with other species.
To observe how cell boundaries change during male tail morphogenesis (e.g. cell fusions
in tail tip and seam cells), use a him-5 (high incidence of males) strain that has ajm-1::gfp
(adherens junction molecule-1, a component of belt adherens junctions in epithelia).
5. Live analysis of sperm behavior. Mate red (histone) sperm males with
hermaphrodites of other strain. Use mCherry::Histone males. Observe the migratory
behavior of the red sperm in the spermatheca. If the hermaphrodite had green sperm (eg
strain TH32), what happens to the red histones after fertilization? Are there histones
retained in the male pronucleus or displaced by green histones from the “mother” worm?
When is the red histone gene from the sperm-derived chromosome transcribed,
translated, and incorporated into chromosomes in blastomeres?
18
6. Ovulation time-lapse. A time-lapse study of ovulation in C. elegans can be
performed by first anesthetizing adult hermaphrodites in a solution of M9 with 0.2%
tricaine and 0.02% levamisole for 30 minutes in a glass depression slide. The
anesthetized worms are then mounted on 5% agarose pads with 10-20 µl of anesthetic
and covered with an 18 mm glass coverslip. Seal the edges of the coverslip with Valap
and capture images at 10-20 second time intervals by DIC optics using a 100X objective.
Try performing time-lapse by also visualizing the sperm containing red-fluorescent
protein-tagged histone H2B.
7. Using photo-conversion to study basement membrane dynamics. Recently, the
photo-switchable fluorophore Dendra2 was developed for use in C. elegans. Based on
principles similar to photo-activatable GFP where a light induced conformational change
in the fluorophore alters the fluorescent properties of the molecule, Dendra2 initially
fluoresces under blue light (green emission, just like GFP), however, after brief exposure
to UV light Dendra2 converts and is thereafter excited by green light (red emission).
Using the laminin::Dendra2 transgenic worms and an appropriate UV laser, you should
be able to photo-convert a region of interest within the basement membrane (for example,
under the anchor cell prior to invasion) and then track the fate of the basement membrane
that was converted from green to red fluorescence. Alternatively, you could photoconvert all of the basement membrane in the animal from green to red fluorescence and
then image the animal at later time points to determine how newly synthesized laminin is
added to a growing basement membrane (the new laminin will be green, whereas
preexisting laminin will remain red).
8. Vulval Development in other species (an evo-devo project): A recent Current
Biology paper (Kiontke et al. 2007) provides a well characterized overview of many of
the developmental changes that have occurred in nematode vulva development across a
wide range of rhabditid species. While many characters have changed multiple times
during the evolution of the rhabditid nematodes, the authors note one character that is
unchanged in all of the species observed:
19
The VulD cell (which arises from the 2˚ lineage (P5.p and P7.p) does not divide.
From Kiontke et al. (2007): Phylogram showing character states of 2˚ vulval lineages in
rhabditid nematodes. Note VulD never divides in all species examined.
Recent evidence from the Sherwood lab suggests that the VulD cells play an
important role in stabilizing the boundary of the gap created in the basement
membrane by the AC during invasion.
The VulD cells (long white arrows)
form the boundary with the overlying
BM (green, LAMININ::GFP; short
white arrow), stabilizing the gap
created following AC invasion into the
vulval epithelium.
One explanation for the invariance of this character is that the VulD cell must remain
undivided to stabilize the overlying basement membrane (BM). To test this hypothesis
you can perform the following experiments:
1) Does the VulD cell form the boundary in non-C. elegans species? Using DIC optics you
can examine early L4 stage worms and identify which vulval cells form the boundary
with the overlying BM. We have mitotracker available if you would like to try to
visualize the BM using fluorescence in other species. Note: mitotracker works better in
some species than in others. See David Matus for a mitotracker BM staining protocol.
20
2) Does laser ablation of VulD at the Pn.p 4-cell stage in C. elegans result in a “sloppy”
boundary? (Perform these experiments in a strain with LAMININ::GFP so you can
visualize the BM following your ablations.)
VI. Lab schedule
Lab 1, Thursday afternoon
General demo introduction (all faculty and TAs)
Set up nematode mating plates
Individual demos:
a) mounting early embryos (Paul/ Amy)
b) mounting and dismounting larvae (Elliott)
c) laser ablation (Dave S)
d) wormbase (Dave M)
Lab 2, Thursday evening
Score RNAi plates for uterine-vulval defects (round 1)
Chalk Talk: Amy Maddox
Overview of non-RNAi experiments (All faculty)
Independent experiments
Lab 3, Friday Afternoon
Independent experiments
Lab 4, Friday Evening
Score RNAi plates for uterine-vulval defects (round 2)
Independent experiments
Friday Evening Lecture Series, 8 o'clock, Lillie Auditorium
Lab 5, Saturday afternoon
Independent experiments
Lab 6, Saturday evening
Chalk Talk: Paul Maddox
Independent experiments
21
V. Appendices
A. Strain list (transgenes are annotated as promoter::PROTEIN)
N2 (Wild-type Caenorhabditis elegans var. Bristol)
Caenorhabditis briggsae
Oscheius tipluae
Oscheius myriophila
Pristonchus pacificus
Panagrolaimus superbus
Acrobeloides butschlii
Cephalobus sp. (DWF1301)
Zeldia sp. (DWF1701)
Panagrellus redivivus (MT8872)
ajm-1::GFP
ceh-22::GFP
elt-2::GFP
edIs20
lin-26::GFP
hlh-1::GFP
cdh-3::GFP
cog-2::GFP
egl-17::GFP
unc-31
him-5
pkd-2::GFP
pkd-2; him-5
pAC::mCherry, laminin-beta::GFP
pAC ::mCherry ::moeABD ; laminin::GFP
laminin::Dendra2
nmy-2:: NMY-2::GFP (JJ1473)
pie-1::GFP::tubulin (AZ235)
pie-1::GFP::H2B (AZ212)
pie-1::GFP::PLCδPH (OD58)
pie-1::GFP::UNC-59 (OD26)
GFP::gamma-tubulin; GFP::Histone (TH32)
OD122
F57B10.1>GFP::beta-tubulin
cdh-3>beta-tubulin::GFP
22
1. Mutant strains
him-5. High incidence of male strain. Approximately 16% of the self-progeny are male.
pkd-2. pkd-2 encodes an ortholog of human PKD2 (OMIM:173910; mutated in
autosomal dominant polycystic kidney disease) that is expressed in the cilia of three types
of male-specific sensory neurons.
2. Strains expressing tissue-specific GFP markers
Strains listed below contain integrated transgenes that direct tissue-specific expression of
the Green Fluorescent Protein (GFP). Many of the GFP transgene strains we have
brought are homozygous for a chromosomally integrated ‘S65C’ variant reporter and are
marked by a dominant ‘roller’ phenotype (animals have a “C” shaped appearance).
Depending upon the nature of the fusion transgene, fluorescence may be cytoplasmic
(e.g. edIs20), localized to subcellular structures (e.g. ajm-1::GFP).
ajm-1::GFP expresses in epithelial adherens junctions, thus outlining all epithelia. This
can be used to study cell fusion, cell movements, etc. ++ (adults); + (embryos and
larvae).
ceh-22::GFP: expresses in differentiated pharynx muscle cells. +++ (all stages). Pharynx
cells are made autonomously from MS and by induction in AB; thus, this marker is useful
for detecting early embryonic interactions.
elt-2::GFP (rrIs1): expressed in E lineage (~4E) and differentiated gut cells (very strong
fluorescence, visible under the fluorescence dissecting microscope). This strain does not
carry the Rol marker. +++ (all stages)
edIs20: This strain carries a fusion of a putative signal transduction gene (F25B3.3) to
GFP. Expression begins in embryonic neural precursors and is later seen in a large part
of the nervous system. In particular, the ventral nerve cord and commissural interneuron
processes are most impressive. The edIs20 strain is in a Him genetic background,
meaning that it generates spontaneous males at high frequency due to X-chromosome
nondisjunction. The male mating structure, the bursa, contains sensory rays that can be
visualized with this marker (very strong fluorescence, visible under the fluorescence
dissecting microscope). +++ (late embryos and older)
lin-26::GFP: expresses in nuclei of hypodermal and other non-neuronal ectodermal cells.
+ (embryos); ++ (later stages)
hlh-1::GFP: expresses in body wall muscle cells. ++ (all stages)
cdh-3::GFP: (syIs50) high levels of expression in the anchor cell and seam cells.
23
cog-2::GFP: Transcriptional reporter for the cog-2 gene, which encodes a sox domain
protein expressed in the uterine pi cells that form the mature uterine-vulval connection.
cog-2 is also expressed faintly in uterine rho cells (see Fig. 3A) and brightly in the body
wall muscles.
egl-17::GFP: Transcriptional reporter for the egl-17 gene, which encodes a fibroblast
growth factor-like protein that is expressed in P6.p and its descendants into the early L4
stage (expressed highest in VulE and VulF cells) before it is shut off. It is then turned
back on in the P6.p descendants VulD and VulC during the mid L4 stage. egl-17::GFP is
also be expressed in the dorsal uterine cells during this time.
nmy-2::GFP: Non-muscle myosin chain. Expressed in the germline and somatic gonad.
pkd-2::GFP: (nIs128) pkd-2 encodes the C. elegans homologue of the human disease
gene polycistin-2. pkd-2::GFP is expressed in the male-specific sensory neurons of the
head (the CEMs), rays and hook (HOB).
Embryonic and germline GFPs
The pie-1 promoter drives expression in the germline: the gonad and embryos. This
fairly strong promoter is routinely used with GFP fusions to study protein behavior in the
early embryo in lieu of identifying and isolating the protein of interest’s own regulatory
sequences.
pie-1::tubulin::GFP: Expressed in germline and early embryos.
pie-1::GFP::H2B: Histone::GFP fusion in germline
OD3 pie-1::tubulin::GFP: Expressed in germline and early embryos.
AZ244 pie-1::GFP::H2B and pie-1::tubulin::GFP: in germline
TH32 pie-1::GFP::H2B and pie-1::gamma-tubulin::GFP: in germline
B. Founder cell lineages and cell fate markers
Consider the following lineage when you design your ablation experiments. The general
experimental outline is to ablate a cell or subset of cells, and then assay the fates of the progeny
of the remaining cells. A variety of fate markers are available. However, we suggest using one
of the GFP-expressing strains. This will allow you to directly assess the results of your ablation
after allowing time for development.
24
Po
P1
P2
EMS
P3
Founder cell
AB
MS
E
P4
D
C
20
32
20
Gut cells
Seam cells 20
Pharynx : structure
muscle
16
2
19
18
Body muscle
1
28
Embryonic deaths
98
14
1
GFP expressing strains. The constructs that cause tissue-specific GFP expression are
particularly useful markers (see the strain list).
Gut granules. The cytoplasm of intestinal cells contains refractile and autofluorescent
"gut granules" ("Rhabditin"). In intact embryos they begin to appear at about the 100150 cell stage (3-4 hours post fertilization). These can be observed under polarized light.
Simple polarized light can be obtained by carefully pulling out the Wollaston prism from
the Nomarski scope. Gut nuclei can be visualized with elt-2::GFP.
Pharynx morphology. Look for the distinct structures of the pharynx tissue. Some
ablations may produce a partial pharynx, such as the posterior bulb. You may want to
refer to pharynx morphology of an intact L1. Pharynx muscle can also be visualized with
ceh-22::GFP.
Muscle twitching. Early in the morphogenesis stage of wild type development (6-7
hours post fertilization), "twitching" movements start. These are indicative of muscle
function, and the embryo soon develops into a squirming larva. If body muscles are
present in partial embryos, they can form functional tissues, and may begin twitching at
about the same developmental time. This activity will subside over time. Muscle cells
can also be visualized with hlh::GFP.
25
C. Sensory organs of the male tail
(Ventral; From the worm atlas; scanning
EM-wrinkled tail)
(DIC image, David Fitch)
Sensory Rays
9 bilateral pairs of sensory rays, numbered 1 (anterior) to 9 (posterior). Each ray has a structural
cell and two sensory neurons (RnA and RnB). Except for ray 6, all neurons have contact with
the outside. The sensory rays are required for response to hermaphrodites dorsally and play a
role in the male's ventral response to hermaphrodites in combination with ventrally located
sensory structures, e.g. hook sensillum, hook and spicules. Rays are also involved in turning
behavior.
The Hook Sensillum
The hook sensillum is composed of 2 sensory neurons (HOA and HOB) and two support cells,
and is closely associated with the hook, a sclerotic structure that is shaped like a hook and
derived from a single cell. This entire structure can be removed by ablation of the epidermal
P10.p cell in the late L2 or early L3 stage. The hook sensillum is required for vulval location.
The Post-cloacal Sensilla (p.c.s)
Each post-cloacal sensilla is made up of 3 sensory neurons--PCA, PCB and PCC--and three
support cells. These sensilla may be involved with precisely positioning the spicule over the
vulval opening.
The Spicules
Each spicule contains 2 sensory neurons, SPD and SPV. The processes of these neurons extend
the length of the spicule and are open to the environment. Each spicule also has a motor neuron,
the SPC. The spicule structure is, not surprisingly, required for sperm transfer. The SPD sensory
neuron and SPC motor neuron are also required for spicule insertion. The SPV sensory neuron
regulates sperm transfer.
26
D. Gamete maturation and oöcyte ovulation
from: McCarter et al., Developmental Biology 205, 111-128 (1999)
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