Microbial diversity and community respiration in freshwater

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Research
Cite this article: Ho¨lker F, Wurzbacher C,
Weißenborn C, Monaghan MT, Holzhauer SIJ,
Premke K. 2015 Microbial diversity and
community respiration in freshwater sediments
influenced by artificial light at night. Phil.
Trans. R. Soc. B 370: 20140130.
http://dx.doi.org/10.1098/rstb.2014.0130
One contribution of 14 to a theme issue
‘The biological impacts of artificial light at
night: from molecules to communities’.
Subject Areas:
environmental science, ecology
Keywords:
DNA metabarcoding, next-generation
sequencing, light pollution, primary
production, carbon turnover
Microbial diversity and community
respiration in freshwater sediments
influenced by artificial light at night
Franz Ho¨lker1, Christian Wurzbacher1,2, Carsten Weißenborn1,
Michael T. Monaghan1,2, Stephanie I. J. Holzhauer1 and Katrin Premke1,3
1
Leibniz Institute of Freshwater Ecology and Inland Fisheries (IGB), Mu¨ggelseedamm 301/310,
Berlin 12587, Germany
2
Berlin Center for Genomics in Biodiversity Research, Ko¨nigin-Luise-Strasse 6– 8, Berlin 14195, Germany
3
Leibniz Centre for Agricultural Landscape Research (ZALF), Eberswalderstrasse 84, Mu¨ncheberg 15374, Germany
An increasing proportion of the Earth’s surface is illuminated at night.
In aquatic ecosystems, artificial light at night (ALAN) may influence microbial
communities living in the sediments. These communities are highly diverse
and play an important role in the global carbon cycle. We combined field
and laboratory experiments using sediments from an agricultural drainage
system to examine how ALAN affects communities and alters carbon mineralization. Two identical light infrastructures were installed parallel to a drainage
ditch before the start of the experiment. DNA metabarcoding indicated
that both sediment communities were similar. After one was lit for five
months (July–December 2012) we observed an increase in photoautotroph
abundance (diatoms, Cyanobacteria) in ALAN-exposed sediments. In laboratory
incubations mimicking summer and winter (six weeks each), communities in
sediments that were exposed to ALAN for 1 year (July 2012–June 2013)
showed less overall seasonal change compared with ALAN-naive sediments.
Nocturnal community respiration was reduced in ALAN-exposed sediments. In long-term exposed summer-sediments, we observed a shift from
negative to positive net ecosystem production. Our results indicate ALAN
may alter sediment microbial communities over time, with implications for ecosystem-level functions. It may thus have the potential to transform inland
waters to nocturnal carbon sinks.
Author for correspondence:
Franz Ho¨lker
e-mail: [email protected]
1. Introduction
Electronic supplementary material is available
at http://dx.doi.org/10.1098/rstb.2014.0130 or
via http://rstb.royalsocietypublishing.org.
Artificial light at night (ALAN) is increasingly recognized as a major component
of global environmental change [1,2]. Through its effects on the behaviour and
physiology of organisms, ALAN is a potentially important driver of ecosystem
dynamics [3–6]. There is increasing evidence that ALAN can trigger physiological and behavioural responses in organisms even at very low light levels (below
1 lux) [5,7], making it an ecologically significant light source despite being only
a small fraction of daytime sunlight.
Light can impact biological systems in different ways. It serves as an information source to organisms that, along with temperature, drives daily and
seasonal cycles of behaviour, phenology and physiological change [5,8]. Light is
also the main source of energy for most primary producers and thus a key factor
influencing the community structure, biomass and productivity of phototrophic
communities [9].
Freshwater ecosystems may be particularly susceptible to changing light
regimes at night. ALAN is found where humans are found, and half of the
human population lives within 3 km of a surface freshwater body [10]. Experiments on the effects of ALAN in aquatic systems to date have demonstrated
physiological and behavioural responses of individual species [3], and community
effects over one [11,12] or several [13] generations. ALAN may have consequences
for aquatic primary producers because photosynthesis can occur at very low light
levels. Minimum light requirements for growth among microalgae are around
& 2015 The Author(s) Published by the Royal Society. All rights reserved.
(b) Field sampling
Benthic microbial communities in the two sites were sampled in
July 2012 (summer) before the start of the artificial light treatment,
and in December 2012 (winter), five months after artificial illumination at the lit site. A sediment corer (Uwitec, Austria; inner
diameter: 60 cm) was used to obtain surface sediment (0–2 cm
depth) and individual samples (three replicates per site) were
placed directly into 50 ml Falcon tubes and stored in the dark at
2208C prior to DNA extraction. For the laboratory incubation
experiment, 28 sediment cores from each site were taken in each
field at the end of June 2013. The first 2 cm of all 28 cores were
pooled in one sterile container per site (totalling 1.6 l sediment
each). Water samples were taken on the same day, filtered (GF/
C-filter, Whatman Ltd, USA), and stored in a climate chamber
until the start of the experiment (compare insets of figure 1a,c,d).
Dissolved oxygen, pH, conductivity, temperature, chlorophyll
(YSI 6600 V2 data probe, YSI, Yellow Springs, USA) and precipitation (Vaisala Weather Transmitter WXT520, Vantaa, Finland)
were measured at both sites over a three-week period before each
of the field samplings in July and December 2012, and again in
June 2013. In addition, water samples were taken in 2013 and 2014
to measure carbon and nutrients (dissolved organic carbon
(DOC), total phosphorous (TP), soluble reactive phosphorus (SRP)
and dissolved inorganic carbon (DIC)) according to standard chemical procedures [23,24]. DOC concentrations were determined with
an organic carbon analyser (Shimadzu, TOC-V CPH, Duisburg,
Germany) as non-purgeable carbon after acidification. DIC was
measured using a nitrogen/carbon analyser (multi N/C 3100,
Jena Analytics). Concentrations of TP and SRP were determined
photometrically by the molybdenum blue method [25] using segmented flow analysis (Skalar Scanþþ, Skalar Analytical B.V.). The
two sites were similar in terms of morphology, catchment characteristics and abiotic parameters (electronic supplementary material,
Table S1) of the ditch. There were small but significant differences
in some chemical parameters between the two sites before the
samples were taken (electronic supplementary material, table S1).
(a) Experimental field site
Sediment cores for both field and laboratory studies were taken
from an agricultural drainage ditch at an experimental site in
Westhavelland Nature Park, Brandenburg, Germany (528690 N,
128460 E). Located approximately 70 km northwest of Berlin, it is
one of the darkest areas of Germany at night and is an official
(c) DNA metabarcoding of sediment microbial
communities
DNA was extracted from all sediment samples (field samples
from July and December 2012 and incubated cores) using a
2
Phil. Trans. R. Soc. B 370: 20140130
2. Material and methods
Dark Sky Reserve (www.darksky.org). The experimental site
consists of two grassland fields that were under the same management regime by the same agricultural business (mown twice
per year, no fertilizer). Identical light infrastructures were installed
in two sites along the drainage ditch in early spring 2012. Each
site contains 12 street lights (20 m apart) arranged in three
rows parallel to the ditch. The ditch itself is approximately 5 m
wide and 50 + 26 cm deep. The first row of lights is located 3 m
from the water’s edge. There is little or no water flow in the ditch
except during strong precipitation events. The main flow direction,
as indicated in the topographic map for this area, is from the control
site to the treatment site over a distance of around 800 m (Blatt 3340NO Ferchesar, 1 : 10 000). The experimental sites are separated by
600 m (Euclidian distance) and a row of trees ensures there is no
influence of the experimental lights on the control site. Luminaires
(Schre´der Saphire 1) were mounted at 4.75 m height. Lights (70 W
high-pressure sodium lamps, 2000 K, 96 lm W21; Osram Vialox
NAV-T Super 4Y, Munich, Germany) were turned on in the lit treatment site on 25 July 2012. Nocturnal light levels ranged from 13.3
to 16.5 lux at the water surface (approx. 0.18 mmol m22 s21) and
6.8–8.5 lux (approx. 0.09 mmol m22 s21) at the sediment surface
(50 cm depth) when samples were collected. Light measurements
in the field and laboratory were taken with an ILT1700 lux meter
(range: 0.00167–1 670 000 lux, International Light Technologies,
Peabody, MA, USA)
1 mmol photons m22 s21 (approx. 50 lux) [14]; however, marine
photolithotroph algae are thought to grow at under 0.01 mmol
photons m22 s21 (approx. 0.5 lux), which is slightly above the
light of a full moon on a clear night (0.005 mmol m22 s21,
approx. 0.3 lux) [15]. Poulin et al. [16] found that night-time
irradiance levels comparable with near shore street lighting
(0.08 mmol m22 s21, approx. 6.6 lux) influenced photophysiological processes in Microcystis aeruginosa, a common
freshwater cyanobacterium.
An important next step is to determine whether ALAN can
influence aquatic microbial community composition (MCC)
over time (i.e. species composition and relative abundance)
and whether the impact extends to emergent functional
traits such as community metabolism and associated carbon
turnover. Microbial communities represent an excellent
system for studying temporal changes in community structure
because of their short generation times relative to plant and
animal communities [17]. One possible effect could be a community shift in favour of species or genotypes that are better
suited to survive and reproduce in nocturnal low-light
environments. This may then affect ecosystem-level functions
such as carbon and nutrient turnover.
Community respiration (CR) and net ecosystem production
(NEP) are two of the most pivotal processes in aquatic ecosystem metabolism. They incorporate biochemical pathways that
make organic carbon molecules and energy available to microorganisms [18] and exhibit a strongly positive relationship with
temperature [19]. As ecosystem metabolism is the primary control on carbon cycling in the biosphere, there is substantial
interest in understanding the controls on aquatic ecosystem
metabolism [20]. Freshwater systems are hot spots for carbon
cycling on the landscape scale and play a significant role in
global and regional carbon cycles [21,22]. Consequently,
understanding the controls on ecosystem metabolism and factors influencing aquatic microbial communities is essential for
predicting carbon cycle response to ongoing environmental
change such as the dramatic increase in ALAN.
Here, we examine the microbial communities in the uppermost layer of freshwater sediments of an agricultural drainage
system in northern Germany. We employed a field experiment approach to compare ALAN-naive and ALAN-exposed
sediments, and a common garden incubation to examine
interactions between ALAN and temperature. We used DNA
metabarcoding to examine bacterial and algal communities
and we measured CR in incubated sediment cores at two different temperatures. Our experimental set-up allowed us to test
three hypotheses, namely (i) that the presence of ALAN
would lead to altered benthic MCC by favouring phototrophic
organisms able to carry out photosynthesis at low light levels;
(ii) that changes in MCC would lead to changes in ecosystem
function, e.g. CR and NEP; and (iii) that seasonal changes in
community structure and function would be less pronounced
because of the potential loss of an important seasonal cue.
Transparent incubation cores (acrylic, inner diameter 53 mm;
height 300 mm) were filled with 100 g of sediment sampled in
June 2013 from either the lit or control site (after around 1 year of
ALAN at the experimental site). Incubation cores with the sediment were gently filled up with the filtered ditch water that was
collected on the same day that sediment was collected. It is likely
that oxygenation, which occurred while sediments were being prepared for the experiment, resulted in a loss of Archaea from these
samples (see §3). For the incubations, 10 cores from each site were
placed into a temperature-controlled plant-growth climate
chamber (KBW-720; Binder, Germany). The upper section of the
chamber was programmed for dark (control) conditions at night
and the lower section was programmed for ALAN (treatment) conditions (four treatments five replicates). The sections were
separated by a dark opaque plastic to prevent light exchange. In
each level, light-cassettes (five compact fluorescent lamps, LT—
T8 30 W COLOURLUX plus, 6500 K, NARVA GmbH, Germany)
were placed 50 cm above the incubation cores and these were
used for daytime illumination (3220 + 284 lux 50 cm below at
the incubation cores, approx. 45 mmol m22 s21). In the treatment
section, four LED strips (6300 K, Barthelme GmbH, Nu¨rnberg,
Germany) were installed on the rear panel and were used for
night-time illumination of the treatment incubation cores (71 +
4 lux, approx. 1.3 mmol m22 s – 1). LED intensity was regulated
over a pulse-width modulator (FG-Elektronik, Germany) with a
potentiometer. LED cycles were controlled by a clock timer (type
SB-1, Unitec, Germany). An aeration system prevented the generation of oxygen gradients. Temperature was monitored with data
loggers (Gemini-Data-Loggers, UK). A minimal temperature difference of 0.28C between upper and lower chambers was noted over
the whole experiment.
(e) Data analysis
Chemical parameters were compared using a two-tailed t-test. Data
were checked for homogeneity of variance (Levene test) and normal
distribution (Shapiro–Wilk test) and transformed (ln or 1.1 to 2)
if needed. Alternatively, a Welch test or a Mann–Whitney U-test
was applied. Tests for each data subset were Bonferroni-corrected
for multiple comparisons (n ¼ 6). In the subsets, we tested for
(i) the differences between the two sediment types, (ii) the differences between the two subsequent temperatures, and (iii) the
differences between the light treatments and the controls. The
significance level for all tests in this study was set to 0.05.
The number of DNA sequence reads varied across all
samples; thus a random subsample of 3650 reads (the size of
our smallest sample) for the field samples and 4254 reads for
the laboratory incubation samples was used to make data comparable. Laboratory incubation sequences consisted of paired
reads and thus first had to be overlapped using MOTHUR [30]
by defining a minimum length of 190 and a minimum overlap
of 25 with a maximum of five mismatches. All sequences (field
and laboratory sequences) were then submitted to the SILVA
NGS pipeline [31] using default settings (www.arb-silva.de/
ngs/, accessed May 2014) for data curation and taxonomic classification. This resulted in a taxon-abundance table. The SILVA
NGS table (number of reads for each taxonomic path, corresponding to an organism group with the maximum taxonomic
depth of 20) was used as input for R. Krona charts [32] were created using the group means. Subsequently, MCC was analysed
by multivariate statistics using the vegan package [33]. Nonclassified species (‘no relatives’) were excluded from all datasets
prior to statistical analysis, which resulted in 1141 taxonomic
paths for the field sites and 2195 taxonomic paths for the laboratory experiment. Taxonomic turnover (bsor; calculated with the
Sorensen index (binary) and the Shannon index) was calculated
using a subsampled matrix with the sequencing read counts.
For the MCC, we analysed the field samples using non-metric
multidimensional scaling (NMDS). The matrix was transformed
3
Phil. Trans. R. Soc. B 370: 20140130
(d) Laboratory incubations
The incubation experiment consisted of two different phases,
designed to mimic summer and winter conditions of the ditch
sediment ecosystems. Starting with the summer conditions, the
temperature was maintained at 20.0 + 0.038C, with a light : dark
cycle of 16.5 L : 7.5 D (daylight from 05.00 to 21.30). Throughout
the night (21.30 to 05.00), control incubation cores were in complete
darkness and treatment incubation cores were lit with LED lights.
The summer experiment lasted six weeks, after which temperature
and photoperiod were linearly reduced over a period of 12 days, to
acclimatize the cores to winter conditions. Winter temperature
was maintained at 6.0 + 0.098C with 7.5 L : 16.5 D (daylight from
08.30 to 16.00). In the treatment chamber, cores were again lit
with LED lights throughout the night (from 16.00 to 08.30).
Water samples were taken at the start of the experiment and at
the end of each incubation period (208C and 68C) to measure
DOC, DIC, TP and SRP.
CR in the sediment cores was measured using a needle-type
oxygen microsensor (Optode, PreSens, Microx TX3, Regensburg,
Germany) after mixing the water column [28,29]. This non-invasive
method was used to measure oxygen concentration through a
septum directly in the water column 1 cm above the sediment of
the experimental cores. This resulted in seven measurements per
core at 208C and 16 measurements per core at 68C. To obtain
values, oxygen concentrations were measured three to six times starting 30 min after hermetically sealing the sediment core. The amount
of consumed oxygen was calculated over time and averaged over
all repeated measurements. Single CRday measurements differed
significantly with the progression of the incubations (ANOVA, at
208C and at 68C, p , 0.001), while CRnight measurements were
independent from the measuring date (ANOVA).
phenol–chloroform/CTAB method [26]. Approximately 350 ml
of sediment was used for each extraction. A mechanical beadbeating step was performed using MMX400 (Retsch GmbH,
Germany) for two rounds of 3 min at a frequency of 28.5 s21.
DNA was quantified using a NanoDrop spectrometer (PEQLAB
Biotechnologie GmbH, Germany) and uniformly diluted to a concentration of 52 ng ml21. One microlitre of this was used to amplify
the V8 region of the ribosomal small subunit marker gene with
universal primers (926F and 1392R) from [27].
Two amplicon libraries were constructed, one for the field
study, which was sequenced with 454 pyrosequencing (GS
Junior Titanium bench top sequencer, Roche, 454 Life Science),
and one for the incubation experiment, which was sequenced
with a MiSeq (Illumina). For the 454 sequencing, barcoded
fusion primers were used with conditions as described previously
[27]. The protocol was modified by using the Herculase II Fusion
DNA Polymerase system (Agilent Technologies) with 0.25%
DMSO with 30 cycles of amplification, and by increasing annealing and denaturation times to 45 s. The master mix and samples
were prepared for amplification on a sterile bench to prevent
foreign DNA contamination. Amplicons were gel-checked, purified, pooled and subjected to emulsion PCR following the steps
of the GS Junior Titanium Series manual for Lib-L amplicon
libraries (Roche, 454 Life Science). Illumina sequencing used the
same native primers (926F and 1392R) without barcodes, no
DMSO in the PCR reaction and an annealing temperature of
508C. PCR products were purified with magnetic beads (MagBio
HighPrep, GC biotech) and the library preparation was carried
out with Nextera XT DNA Sample Prep Kit (Illumina) in combination with the 96 Nextera XT Index Kit (Illumina) following the
manufacturer’s instructions. During the preparation, the amplicons are shortened by a transposase. The resulting sequencing
library was subjected to sequencing with the two-times 300 bp
reads (V3 chemistry, Illumina).
(a)
(b)
4
stress = 0.09
0.5
C
0.4
Dec-T
Jul-T
0.1
0.2
0.3
0
0.1
0.2
0.3
0.1
Jul-C
17.5% of variance explained
(c)
(d)
stress = 0.18
stress = 0.18
Shannon
T
C
temperature
y
CR da
ni
gh
light
axis-1 - seasonality/temperature
20°C ALAN
20°C night
6°C ALAN
DOC
CR
t
axis-1 - seasonality/temperature
6°C night
night
ALAN
CRnight
Figure 1. MCC presented as NMDS (axis 1 was always aligned with the seasonal shift/temperature) and as partial RDA in (a) field sediments and (b–d) incubated cores. Ellipses
indicate the standard error around the centroid for each core type (origin and experimental treatment). Insets are Krona charts with two taxonomic levels of the microbial
community displayed at the start of each experiment (red, Bacteria; brown, Eukarya; blue, Archaea; green, no relatives; C, ALAN-naive (control) sediment; T, ALAN-exposed
(treatment) sediment). (a) Ditch sediment samples were analysed using a weighted presence/absence matrix and Euclidean distances, calculated and presented as NMDS.
(b) Laboratory-incubated cores were examined using partial RDA of the sediment community. Sediment type explained 6.4% of the variation. The residuals were constrained
with temperature and the mean night respiration. The RDA model was significant for both terms (ANOVA, temperature p , 0.001; CRnight p , 0.05, 2000 permutations).
(c,d) NMDS ordinations of laboratory-incubated cores (analogous to (a)) with the two sediment sources (c: ALAN-exposed; d: control). The arrows mark significant parameters
which were fitted into the ordination by R (envfit, p , 0.05). (Online version in colour.)
into a probability presence/absence matrix (function ‘drarefy’
with the smallest number of sequences) and served as the basis
for calculating the NMDS ordination by applying Euclidean distances (figure 1a). The ordination was rotated so that the first axis
corresponds to season. The separation of the sampling dates and
sites was visualized by drawing standard-error ellipses around
group centroids with a confidence level of 0.95. This was supported by a permutational multivariate analysis of variance
(PERMANOVA). Seasonal differences in major taxonomic
groups between the ALAN-naive and ALAN-exposed areas
(figure 2a) were examined by subtracting each December value
from each July value. We removed all minor microbial groups. Significance was tested with a Mann– Whitney U-test and was not
corrected for multiple comparisons.
We analysed the samples from the laboratory set-up in a similar
way. We started the analysis of the MCC by testing the possible
influence of temperature and CRnight on the MCC. This was done
by calculating a partial constrained redundancy analysis (RDA)
based on the original quantitative matrix (figure 1b). The effect of
the two sediment types was thereby removed. Afterwards, the
CRnight was fitted to the RDA by using the function ‘ordisurf’,
which applies a generalized model. We further examined the qualitative structure of the MCC as described for the field samples (figure
1c,d). The two NMDS ordinations significantly underlying environmental parameters were explored by using the vegan function
‘envfit’ with 2000 permutations (verified by a Spearman-based
Mantel-test), while the separation of light treatment and temperature
was visualized by drawing standard-error ellipses around centroids
with a confidence level of 0.95 accompanied by a PERMANOVA.
Changes in the major taxonomic groups (figure 2b) were
measured by subtracting the 208C values from the starting mean
values of each sediment type and by subtracting the 68C values
from the mean values of the 208C incubations. Significance was
tested with a Mann–Whitney U-test (not corrected for multiple
Phil. Trans. R. Soc. B 370: 20140130
Dec-C
3.1% of variance explained
T
(a)
5
*
2.8
*
*
*
*
*
0
*
*
*
treatment field
*
control field
–2.8
*
(b)
control ALAN
*
*
*
*
0
control night
*
*
*
*
*
*
*
treatment ALAN
treatment night
photoautotrophs
–2
Eukaryota; SAR; Stramenopiles
Eukaryota; SAR; Dinophyceae
Eukaryota; SAR; Ciliophora
Eukaryota; Opisthokonta; Holozoa
Eukaryota; Cryptophyceae
Eukaryota; Archaeplastida
Bacteria; Verrucomicrobia
Bacteria; Spirochaetae
Bacteria; Nitrospirae
Bacteria; Lentisphaerae
Bacteria; Gammaproteobacteria
Bacteria; Firmicutes
Bacteria; Deltaproteobacteria
Bacteria; Cyanobacteria
Bacteria; Chloroflexi
Bacteria; Chlorobi
Bacteria; Candidate division OP3
Bacteria; Betaproteobacteria
Bacteria; Bacteroidetes
Bacteria; Alphaproteobacteria
Archaea; Euryarchaeota
Figure 2. Changes of the inferred relative abundance in selected microbial groups in (a) sediment samples between July and December 2012; and (b) incubated
cores during the laboratory experiment between the end of the 208C incubations and the end of the 68C incubations. Positive changes indicate an increase towards
December. Significant differences between the ALAN-naive and ALAN-exposed sediments are marked with asterisks (Mann – Whitney U-test, p , 0.05). For
Stramenopiles/diatoms only autotrophic subgroups are considered, Chloroflexi are here considered to be heterotrophic since the subgroups were exclusively
heterotrophic (see electronic supplementary material, figure S1).
comparisons). Last, we employed a permutation analysis on the
laboratory incubation data for identifying single taxonomic
groups at the deepest level that was resolved. This was written
as a function in R, which randomly subsampled the community
matrix 500 times and performed a Mann –Whitney U-test per taxonomic group for each subsampling in order to cope with the type I
error caused by the subsampling. The Pq-values show the fraction
of non-significant Mann–Whitney U-tests in 500 random subsamples. The significance level for the Pq-value was set to 0.05
without additional correction for multiple comparisons.
3. Results
(a) Field experiment
The microbial communities (Bacteria, Archaea and Eukarya)
in the freshwater sediments of the two ditch sites (ALANnaive and ALAN-exposed) showed a large degree of overlap
in their overall structure at the start of the study in July 2012
(figure 1a, inset) and in taxonomic composition (bsor ¼ 0.339
between ditches; bsor ¼ 0.321 between replicates within each
site; electronic supplementary material, figure S1). After five
months of illumination, the communities at the two sites
remained similar (PERMANOVA, p . 0.05) although the
NMDS ordination on the two sampling dates detected a significant difference between the two sites in December 2012
(figure 1a). There was a significant seasonal trend (PERMANOVA, p , 0.01), with seasonal difference appearing to be
smaller at the ALAN-treated site.
From the taxon list generated from DNA metabarcodes, a
number of microbial groups were present that could potentially respond to artificial illumination (figure 2a). Of these,
there was an increase in the relative abundance of photoautotrophic Stramenopiles (dominated by diatoms in our
samples, see electronic supplementary material, figure S1)
in the ALAN-exposed sediments five months after lights
had been switched on. Mixotrophic organisms (Cryptophyceae
and Dinophyceae) were either unaffected or were reduced after
five months of illumination. Some of the major heterotrophic
microbial groups (Bacteroidetes and Deltaproteobacteria) also
were reduced after illumination.
(b) Laboratory incubation experiment
Starting conditions in the sediments from the two sites (ALANnaive and ALAN-exposed) differed slightly but consistently
in DOC concentrations (ALAN-exposed . ALAN-naive,
mean DDOC ¼ 2.7 mg l21, mean DOC ¼ 23.3 + 3.1 mg l21).
Concentrations of DIC, TP and SRP were similar. Mean SRP concentrations were 4.9 + 2.8 mg l21. Following the experiment,
cores exhibited significant differences between the two temperature regimes in all measured parameters (table 1). Microbial
Phil. Trans. R. Soc. B 370: 20140130
*
2
Table 1. Significant differences (Bonferroni-corrected) between the two temperature regimes in all measured parameters in sediments from the two sites in the
agricultural drainage ditch that were used for the experimental incubation. n.s., non-significant.
DOC
DIC
CRday
seasonal effect (208C versus 68C)
TP
all data
2088 C
688C
effect in sediment incubations pre-exposed to ALAN
*
n.s.
***
*a
**
n.s.
higher
lower
n.s.
all data
*
no ALAN
n.s.
ALAN
lower
effect in 208C incubations
***
n.s.
higher
DIC
DOC
***
***
***
**
n.s.
***
lower
higher
CRday
CRnight
***b
n.s.
***
**
***b
n.s.
lower
higher
all data
n.s.
208C
n.s.
68C
**
effect in ALAN incubations
lower
*a
n.s.
n.s.
lower
ALAN—control site
DIC
all data
*
208C
n.s.
68C
**
lower
ALAN—pre-exposed site
all parameters
all data
n.s.
208C
n.s.
68C
n.s.
n.a.
ALAN—all data
DIC
CRnight
*p , 0.05; **p , 0.01; ***p , 0.001.
a
Mann – Whitney U-test.
b
Welch test.
communities at the start of the experiment were similar in the
two sites (bsor ¼ 0.388 between sites; bsor ¼ 0.385 between replicates within each site; see insets of figure 1c,d). Differences
primarily were the result of differing abundance (i.e. relative
number of sequencing reads) at the lowest taxonomic level (electronic supplementary material, figure S1 and table S2). The
microbial communities from all incubations were structured significantly by temperature (68C and 208C, partial RDA, p , 0.001)
and respiration (CRnight; partial RDA, p , 0.05; figure 1b). Qualitative MCC of each sediment type revealed that the ALANexposed sediment community showed no group separation at
all (PERMANOVA, p . 0.05), i.e. no seasonal response to the
temperature trigger or a light response.
In the ordinate (figure 1c), a weak effect of ALAN was visible,
which corresponded to a decreased Shannon index (Mantel test,
r ¼ 0.163, p , 0.01). This was in strong contrast to the ALANnaive sediment MCC, which exhibited a significant group separation (PERMANOVA, p , 0.01). This could be attributed to a
pronounced seasonal effect and absence of a light effect, as
above (PERMANOVA, p , 0.001 and p . 0.05, respectively). In
the corresponding ordinate (figure 1d), the seasonal effect was
apparent, triggered by a correlation of the MCC with seasonal
variables (temperature and DOC) together with influences
from the CRday/night (Mantel test, r ¼ 0.473, p , 0.001).
Changes in relative abundance of responsive taxa again
included a potential increase in diatoms with light (Stramenopiles in figure 2b). Cyanobacteria also exhibited a
pronounced response in cores from the ALAN-exposed site.
The communities in the ALAN-naive sediment behaved similarly to the field data, i.e. the sediment communities showed
clear seasonal differences (figure 2a). At the lowest available
taxonomic level, several photoautotrophic (Cyanobacteria,
green algae) and heterotrophic organisms were affected by
light in the ALAN-exposed sediment over the course of the
experiment (table 2). In subsets of the data (i.e. at the 208C
incubations or the 68C incubations), single photoautotrophic
groups increased in abundance (208C, diatom Nitzschia;
68C, Cyanobacteria; see detailed electronic supplementary
material, table S2). We found no significant changes to taxonomic groups in the ALAN-naive sediment. Only in subsets
of the ALAN-naive data (i.e. at the 208C incubations or
the 68C incubations) could single non-photoautotrophic taxonomic groups be identified (see detailed table in electronic
supplementary material, table S2).
The exposure to ALAN had a clear effect on the CR at night
(CRnight) (table 1). The CR during the day (CRday) was lower
under summer conditions for the ALAN-naive sediment
(figure 3b, CD (ALAN-naive sediment, dark at night, day
measurements), and table 1). When compared directly,
median CRnight was lower across all light-treated incubations
except for the ALAN-naive sediment community at 208C
(figure 3b, CLN (ALAN-naive sediment, lit at night, night
measurements)). This lower CR under ALAN conditions
points to oxygen production by photosynthesis, especially in
sediment communities at summer conditions previously
exposed to ALAN (figure 3d, TLN (ALAN-exposed sediment,
lit at night, night measurements)).
4. Discussion
Artificial illumination near surface freshwaters has the potential
to be a viable night-time energy source for microbial phototrophs, some of which can carry out photosynthesis at light
levels that are near to those generated by ALAN. This could
result in changes to the microbial community as conditions
may favour taxa that benefit from nocturnal (low-level) light.
These changes could, in turn, influence ecosystem-level processes such as carbon turnover. Such changes may also vary
Phil. Trans. R. Soc. B 370: 20140130
**
a
site differences
6
(a)
(b)
0.8
0.6
0.2
*
0.1
*
R
*
0
0
*
–0.2
P
–0.4
–0.6
–0.8
–0.3
CL
CD
CLN
CDN
CL
CD
CLN
(d)
0.8
0.3
*
*
0.2
0.6
*
CDN
*
0.4
0.1
0.2
0
0
–0.2
–0.1
–0.4
–0.2
–0.6
–0.8
–0.3
TL
TD
TLN
TDN
TL
TD
TLN
TDN
21 21
Figure 3. CR (mg O2 L h ) of the incubated sediment cores during the day (open box plots) and at night (shaded box plots) at 68C (a,c) and 208C (b,d ). X-axis
labels indicate sediment source (T, sediment from ALAN-exposed (treatment) ditch; C, sediment from ALAN-naive (control) ditch) and incubation treatment (L, lit at
night; D, dark at night). N, night measurements. The horizontal line indicates the transition between gross ecosystem production (P) and respiration (R). Box plots
(median, 25th percentile, 90th percentile, outliers) depict means of measurements made over the six-week period of each experiment. Asterisks denote significant
difference between control and treatment conditions. (Online version in colour.)
Table 2. Significant changes in relative abundance of selected taxa in incubated cores following illumination at night in sediments pre-exposed to ALAN during
the whole course of the experiment (both temperature regimes). Significance is based on a permutated Mann– Whitney U-test ( p , 0.05). Pq-values represent
the proportion of negative Mann –Whitney U-tests. P, photoautotroph; M, methylotroph; H, heterotroph.
domain
ID
taxon
trend
Pq
function
comments
bacteria
358
candidate division TM7
2
*
H
filamentous, stabilize biofilms
449
455
Cyanobacteria, Snowella
Cyanobacteria, Subsection III, Family I
þ
þ
*
***
P
P
psychrophilic
488
Cyanobacteria, MLE1– 12
þ
**
P
612
802
Firmicutes, Ruminococcaceae, Incertae sedis
Alphaproteobacteria, Hyphomicrobium
2
2
*
*
H
M
1552
1597
Chloroplastida, Charophyta, Mesotaenium
Chloroplastida, Chlorophyta, Scenedesmus
þ
2
**
*
P
P
1814
Opisthokonta, Nucletmycea, LKM11
2
***
H
algal decomposer/parasite
1874
Alveolata, Ciliophora, Didinium
2
***
H
feeds on other ciliates
eukaryota
isolated from rumen
some photo-heterotrophy
*pq , 0.05; **pq , 0.01; ***pq , 0.001.
temporally because day length and light levels (along with
temperature) act as seasonal cues for many organisms in natural
systems. We carried out field and laboratory experiments to
examine changes to microbial community structure and
carbon mineralization in sediments of an agricultural drainage
system. The field experiment started when artificial lights were
Phil. Trans. R. Soc. B 370: 20140130
–0.2
community respiration (mg O2 L–1 h–1)
0.4
0.2
–0.1
(c)
7
community respiration (mg O2 L–1 h–1)
0.3
8
?
CO2
CO2
NEP < CR
summer
NEP ≥ CR
summer
NEP < CR
NEP < CR
Figure 4. Summary of long-term effects of ALAN on the microbial community in aquatic sediments. Although there is a general decrease of diversity (Shannon
index) with ALAN, there is an increase in photoautotrophs (green/grey symbols, photoautotrophs; white symbols, heterotrophs). The ALAN-exposed communities also
show less overall seasonal change in community structure between summer (red dotted rectangle) and winter (blue rectangle). This has implications for ecosystemlevel functions such as carbon mineralization and could thus shift the system from negative to positive NEP. (Online version in colour.)
switched on, and control and treatment communities were
compared after five months of exposure. The laboratory experiment sought to verify some of the findings from the field
and examine how community changes in ALAN-exposed (for
1 year) and ALAN-naive communities were linked to CR.
Such a long-term approach, including multiple generations of
most microbes, potentially provides an important insight into
temporal change in community structure and associated process rates arising from light. This provides an important
advance over most previous ALAN studies which focused on
describing ALAN-exposed communities [34] or examining
short-term effects on ALAN-naive communities [11,12].
We observed increased abundance in some phototrophic
taxa in sediments after five months of illumination. A pronounced increase in the abundance of diatoms suggests that
ALAN was used as a source of energy, thus supporting our
first hypothesis. Because nutrients, herbivory, temperature
and other factors combine with light to influence the diatom
communities [35], we further examined changes under controlled laboratory conditions. Phototrophs (diatoms, Cyanobacteria,
green algae) again showed an increase in response to ALAN,
but this was much more pronounced in the pre-exposed site.
Collectively, these results suggest that an illumination period
of over three months induced a weak response and that illumination for 1 year was sufficiently long to prompt a clear change in
what had been an ALAN-naive freshwater microbial community. These data provide a starting point for future studies on
rates of community change.
Several mechanisms might play a role in the response
of Cyanobacteria, diatoms (e.g. Nitzschia) and green algae (e.g.
Mesotaenium) to light at night. Some Cyanobacteria and diatoms
have light compensation points of growth and photosynthesis
below or close to our experimental ALAN level [14,36] and
may thus be able to use ALAN. Others have higher maintenance energy demands and may be unable to use these low
intensities. Some dinoflagellates can photosynthesize and
grow at low photon flux densities but cannot tolerate relatively
high light environments [14]. Thus, the ability to use a large
range of light intensities might be beneficial in shallow aquatic
systems that receive high levels of sunlight and low levels of
ALAN. State transitions, which are a short-term acclimation
mechanism allowing a cell to better balance the distribution
of energy between photosystems I and II, might be beneficial
for Cyanobacteria and diatoms to increase the efficiency of
utilization of absorbed light energy under ALAN conditions
close to our experimental ALAN level [37]. Finally, there is
some evidence that intermittent near-dark periods influence
photoacclimation to photoperiod in a species-specific way [38].
An important objective of our study was to test whether
changes to the communities were accompanied by functional
changes, namely to the process of carbon mineralization. In
the experiment, we observed lower total CR in ALANexposed incubations, which supports our second hypothesis.
It is not clear for all treatments whether this is indicative of
greater primary production, but nonetheless indicates a significant link between the MCC and carbon turnover. Given
the relatively low-light intensities of ALAN compared with
sunlight, a shift to positive NEP might not be expected.
Instead, indirect effects of ALAN, including on animal
(e.g. herbivore) behaviour and plant phenology, could be a
mechanism influencing primary production. However, in
the treatment with ALAN-exposed sediment communities
at 208C, the majority of cores changed to overall autotrophy
(negative respiration) at a level similar to that observed
during the day. In this way, ALAN could increase NEP,
which has implications for ecosystem function.
Night-time irradiance levels in the laboratory (71 lux,
approx. 1.3 mmol m22 s – 1) were brighter than in our field
study, but fell within the higher range of light levels reported
for street lighting. Perkin et al. [12] measured surface light
levels between 0.008 and 4 lux at selected urban stream
locations. Meyer & Sullivan [13] reported light levels ranging
from 8 to 12 lux at open or partly canopied streams adjacent to
‘highly’ lit areas. Hale et al. [39] estimated that 33% of the city
of Birmingham (UK) is covered by natural areas of which 13%
of the area contributed to the total city area with ALAN levels
above 30 lux. The highest known surface light level, 150 lux
(approx. 2.7 mmol m22 s – 1), due to street light was reported
for a small town close to the experimental field site [40]. This
suggests that the experimental set-up mimicked a high but
Phil. Trans. R. Soc. B 370: 20140130
winter
winter
Perkin et al. [12] suggested that artificial light might be
an example of a typical ‘press’ driver [46] that elicits a
‘ramped’ or delayed and slowly increasing response from
many organisms. It is thought that, over time scales of a few
years, microbial composition usually differs from that of undisturbed communities [47]. To reliably predict the ecological
consequences of ALAN in natural systems, it is critical to
have a better understanding of longer term processes that
moderate the susceptibility of communities to an illuminated
environment [48].
ALAN induced shifts in microbial community structure and
ecosystem function and may lead to overall shifts in NEP. As
ALAN increases in global extent [2], increased illumination
of water bodies in rural areas could lead to diminished diurnal
fluctuations and subsequently a shift from negative to positive
NEP. In short, ALAN has the potential to transform inland
waters to a carbon sink during the night. Additional studies
are needed to verify whether this is the case in other aquatic
systems. This information could greatly enhance our ability
to predict system responses to global change.
Data accessibility. DNA sequences have been submitted to the European
Nucleotide Archive (ENA) under the study accession number
PRJEB8381.
Acknowledgements. This is publication 12 of the Berlin Center for
Genomics in Biodiversity Research (http://www.begendiv.de). We
are grateful to Susan Mbedi and Stefan Heller for their help in the
field and laboratory, to Katrin Attermeyer for designing figure 4
and to Tom Shatwell for helpful comments.
Author contributions. F.H., M.T.M., C.W. and K.P. conceived the study.
Ca.W. carried out the respiration measurements. Ca.W. and S.H. carried out the sediment sampling. Ca.W. and C.W. carried out the
DNA metabarcoding. C.W. analysed the sequence data and carried
out all community analyses. C.W., F.H. and K.P. analysed respiration
data. All authors contributed to writing the manuscript.
Funding statement. Research was supported by the Verlust der Nacht
project, funded by the Federal Ministry of Education and Research,
Germany, BMBF-033L038A.
Conflict of interests. We have no competing interests.
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Microbial diversity and community respiration in freshwater

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