Integrative and Comparative Biology
Integrative and Comparative Biology, volume 52, number 5, pp. 636–647
doi:10.1093/icb/ics086
Society for Integrative and Comparative Biology
SYMPOSIUM
Latitudinal Variation in Protein Expression After Heat Stress in the
Salt Marsh Mussel Geukensia demissa
Peter A. Fields,1 Kelly M. Cox and Kelly R. Karch
Biology Department, P.O. Box 3003, Franklin & Marshall College, Lancaster, PA 17604-3003, USA
From the symposium ‘‘Comparative Proteomics of Environmental and Pollution Stress’’ presented at the annual meeting
of the Society for Integrative and Comparative Biology, January 3–7, 2012 at Charleston, South Carolina.
1
E-mail: [email protected]
Synopsis Individuals of a broadly distributed species often experience significantly different environmental conditions
depending on location. For example, the mussel Geukensia demissa occurs intertidally from the Gulf of St. Lawrence to
central Florida; within this range, northern populations are exposed to temperatures cold enough to freeze the tissue,
whereas southern populations can experience temperatures approaching the species’ upper lethal limit. Thus, G. demissa
provides an ideal system with which to study physiological variation in conspecifics occurring across a broad latitudinal
range. We collected G. demissa at five sites from Maine to Florida, encompassing a range of 1900 km, and have used a
proteomic approach to describe how protein expression varies in individuals from the different locations. We acclimated
individuals from each site to common conditions (188C) for 4 weeks, and exposed a subset of these to acute heat stress
(408C). We separated gill proteins using two-dimensional gel electrophoresis and quantified abundances of the resulting
protein spots. Among mussels acclimated to 188C protein, expression profiles were more similar among individuals from
the same site than among sites, but there was no discernible correlation with latitude. In contrast, after acute heat stress,
protein expression among mussels from different locations varied substantially, with 31 of 448 proteins changing in
abundance in the northernmost (Maine) group, compared with 5–11 proteins in the four southern groups. Identification
of 27 of these proteins revealed five functional clusters: chaperones, cytoskeletal proteins, oxidative stress proteins,
regulatory proteins, and a translation initiation factor. Across these functional categories, the two northernmost
groups, Maine and New York, showed the greatest number of proteins that changed significantly in abundance, as
well as the greatest fold-change in abundance for many of the proteins. We conclude that the northern populations
of G. demissa are physiologically distinct from the southern groups, and that the differences in protein-expression profiles
are consistent with greater sensitivity to heat stress to the north.
Introduction
Widely distributed species necessarily experience a
variety of environmental conditions at different locations across their ranges, and thus the types and
magnitudes of physiological stresses experienced by
individuals in these locations can vary significantly
(Somero 2005; Osovitz and Hofmann 2007).
Intertidal organisms in particular have been the subjects of numerous studies relating habitat, physiological stress, and limits of biogeographic range, with
temperature often cited as a significant factor limiting latitudinal distributions (e.g., Hutchins 1947;
Wethey 1983; Bertness et al. 1999; Sorte and
Hofmann 2005; Jones et al. 2010). For example,
there is considerable evidence that many intertidal
invertebrates are exposed to temperatures approaching their thermal maxima, and leading to significant
cellular damage, during emersion (Hofmann and
Somero 1995; Tomanek and Somero 1999; Stillman
2003; Somero 2005). Thus, although body temperature of intertidal organisms does not always closely
match local temperatures of air or water (Gilman
et al. 2006; Helmuth et al. 2009), and temperature
extremes in the intertidal zone do not vary smoothly
in correlation with latitude (Helmuth et al. 2002),
emersion temperatures appear to play a significant
Advanced Access publication May 28, 2012
ß The Author 2012. Published by Oxford University Press on behalf of the Society for Integrative and Comparative Biology. All rights reserved.
For permissions please email: [email protected].
Latitudinal variation in Geukensia demissa protein expression
637
role in setting distribution limits of intertidal organisms. Despite the importance of temperature in helping define biogeographic ranges, however, there is
little information regarding the extent to which localized adaptation, in contrast to phenotypic plasticity, allows survival at different locations along a
broad latitudinal distribution.
Geukensia demissa, the Atlantic ribbed mussel
common to salt marshes along the eastern coast of
North America, is an ideal organism with which to
examine physiological variation across a wide geographic range. It is found in the high intertidal
zone wherever salt marshes occur, and has a
north–south range of 2000 km from the Gulf of
St. Lawrence to northern Florida (Gosner 1978). In
salt marshes, G. demissa is an abundant and ecologically important species, occurring at high density
(over 1000 individuals/m2 in some areas; Bertness
and Grosholz 1985) and associating with smooth
cordgrass Spartina alterniflora. The mussel’s byssus
forms a network with cordgrass rhizomes and
stems, stabilizing the mudflat against erosion, allowing accretion of more mudflat area over time as the
mussel/cordgrass distribution spreads (Bertness
1984), and providing habitat for scores of invertebrate species.
In this environment, G. demissa is exposed to a
remarkable range of environmental stresses that vary
with vertical position, location in the estuary, and
latitudinal position along the coast. For example,
studies have illustrated the species’ ability to tolerate
extreme temperatures, 458C in southern locations
(Jost and Helmuth 2008) and 138C to 228C in
northern locations (Kanwisher 1955); a wide range of
salinities, from 5 ppt (Lent 1969) to 70 ppt (Wells
1961); and significant desiccation, up to 38% of soft
tissue water content (Lent 1968).
We have taken a comparative proteomic approach
to determine the extent to which individuals of
G. demissa collected from across the species’ latitudinal range differ in protein-expression profiles
(PEPs), and what types of proteins vary among mussels from different locations. Proteomic analyses, by
examining expression of hundreds of proteins simultaneously (Gorg 2004), allows a rapid assessment of
cellular responses to environmental stress (Tomanek
2011), including changes in attributes as disparate as
energy metabolism, cytoskeletal elements, molecular
chaperones, and synthesis and degradation of proteins. We addressed the following questions: (1) Do
mussels from widely separated locations maintain
differences in their PEPs after acclimation to
common conditions? (2) Do differences in PEPs
vary in correlation with latitude? (3) Do PEPs of
mussels from different sites change significantly
after acute heat stress (HS)? and (4) If there is a
difference in PEPs after HS, what proteins vary
most significantly? We found that although there is
no clear correlation between latitude and protein expression in unstressed G. demissa, after HS mussels
from the northernmost site exhibited the greatest
proteomic response, both in number of proteins
whose expression pattern changed, and the magnitude of change in abundance of those proteins.
Furthermore, the identity of proteins up-regulated
in northern G. demissa, dominated by chaperone
and cytoskeletal proteins, indicated that mussels
from the northern region of the species’ range are
more susceptible to HS than are their southern
conspecifics.
Materials and methods
Collection and acclimation of specimens
Geukensia demissa were collected from five locations
along the eastern coast of the United States, encompassing much of the species’ latitudinal range (see
Table 1 for a description of collection sites).
Animals were chilled and transported to Franklin
and Marshall College, where they were kept immersed in artificial sea water (ASW; 33–35 ppt;
Instant Ocean, Marineland Labs, Mentor, OH,
USA) in re-circulating aquaria at 18 þ/ 0.28C.
Mussels were allowed to acclimate for 4 weeks
before acute exposure to heat, and were fed 0.1 ml
per individual Instant Algae Shellfish Diet 1800
(Reed Mariculture, Campbell, CA, USA) three
times per week, an amount we have found to allow
long-term maintenance of mussels in the laboratory.
Replicate groups consisted of six control and six
treatment mussels from each location, numbers
chosen to allow simultaneous electrophoresis of 12
two-dimensional gels at one time.
Acute exposure to heat
Six mussels from each location were acutely exposed
to 408C, a sub-lethal temperature that previously we
had found to induce changes in protein expression in
G. demissa. In order to accurately mimic HS as experienced by mussels in the field, animals were
emersed during exposure (Jost and Helmuth 2008).
Each group of six was split between two glass jars
containing 4 cm of sand saturated with ASW, to
avoid desiccation. Jars were capped and immersed
in pre-heated aquaria, with water temperature controlled so that mussels reached the target body temperature (18C) in 25 min. We monitored body
temperature of mussels by drilling a small hole in
638
P. A. Fields et al.
Table 1 Characteristics of the five locations from which G. demissa were collected
Site
Code
Latitude
Longitude
February Tavg/mina (8C)
August Tavg/maxa (8C)
St. Augustine, FL
FL
29.764
81.262
55/28
266/363
Beaufort, NC
NC
34.723
76.677
1
263/324
Chincoteague, VA
VA
37.934
75.419
56/95
246/323
Hewlett Bay, NY
NY
40.622
73.667
15/133
223/337
Damariscotta, ME
ME
44.034
69.532
19/197
32/51
a
Values represent average and minimum/maximum air temperatures from 1984 to 2008 (with the exception of ME, 1986–2008), measured at
coastal-marine automated network (C-MAN) stations operated by the National Buoy Data Center, National Oceanographic and Atmospheric
Administration (www.nodc.noaa.gov/buoy/). The stations selected are those closest to the collection sites, and are intended to indicate general
climatic trends rather than to predict mussels’ body temperatures.
one valve of one mussel per jar, 1 cm from the
posterior edge, and inserting a K-type thermocouple
wire into the mantle cavity; the wire was held in
place by cyanoacrylate glue. There was no significant
difference in protein expression between heatexposed mussels with holes drilled in their shells
versus those with no holes (data not shown).
Target body temperature was maintained for 1 h,
and mussels were returned to aquaria at a temperature of 188C to recover for 24 h before sacrifice. This
recovery period is necessary to allow a robust
protein-expression response to HS. A separate
group of six mussels from each location acted as a
control, and were kept immersed at 188C continually
until sacrifice. No mortality resulted from HS; gill
protein was purified from 60 mussels.
Extraction and purification of protein
Unless otherwise specified, chemicals were purchased
from Sigma Chemical Co., St. Louis, MO, USA. Gills
were excised from controls and treated individuals,
and immediately homogenized (1:4 w/v) in denaturing homogenization buffer [7 M urea (Biorad,
Hercules, CA, USA), 2 M thiourea, 1% ASB-14
(Calbiochem, San Diego, CA, USA), 40 mM Tris
base, 0.5% ampholyte solution (pH 4–7, GE
Healthcare, Piscataway, NJ, USA), 1.2% Destreak
Solution (GE Healthcare), and 0.001% bromophenol
blue)
using
ground
glass
homogenizers.
Homogenates sat (RT) for 1 h to solubilize proteins,
and were centrifuged (16,000g; 30 min; RT) to pellet
insoluble material.
Proteins were precipitated from homogenate supernatants using 10% trichloroacetic acid in acetone
(1:4 v/v supernatant:TCA/acetone). After incubating
(208C) overnight, precipitated proteins were pelleted by centrifugation (16,000g; 30 min, 48C), supernatant was removed, and pellets were washed three
times by vortex-mixing with ice-cold acetone.
Protein was resuspended using a volume of
rehydration buffer (7 M urea, 2 M thiourea, 2% w/v
CHAPS, 2% NP40 substitute, 0.6% pH 4–7 IPG
buffer, 1.2% Destreak Solution, and 0.01% bromophenol blue) equal to the original homogenization
buffer volume. Protein was vortex-mixed and resolubilized overnight at 48C. Samples were centrifuged
(16,000g; 30 min; RT) and supernatant was used immediately for electrophoresis or stored at 808C.
Two-dimensional gel electrophoresis
Proteins were separated by isoelectric point in the
first dimension and by molecular mass in the
second. Protein concentrations were determined
using the 2D Quant kit (GE Healthcare). A total of
200 mL of 1 mg/ml protein were loaded into a single
well of an 11 cm isoelectric focusing (IEF) tray
(Biorad IEF Cell). An immobilized pH gradient
(IPG) IEF strip (11 cm; pH 4–7; Biorad) was laid
over each sample, and covered with mineral oil to
prevent evaporation. Strips rehydrated passively for
6 h and actively (50 V) for 12 h; water-saturated
wicks were placed between the electrodes and the
strips before focusing (500 V for 1 h, 1000 V for
1 h, 5000 V for 1 h, 8000 V for 3 h; all voltage changes
occurred in rapid mode; maximum current 50 mA).
After focusing, oil was blotted from the strips and
they were either frozen (808C) or used immediately
for second-dimension electrophoresis.
In the second dimension, proteins were separated
by mass using precast sodium dodecyl sulfate (SDS)–
polyacrylamide gradient gels (10.5–14%; Biorad
Criterion). Proteins in the IPG strips were equilibrated with SDS, and cysteinyl residues were covalently capped with iodoacetamide to prohibit
formation of disulfide bridges, by 15 min agitation
in equilibration buffer [6 M urea, 375 mM Tris–
HCl pH 8.8, 30% glycerol, 2% SDS (Biorad), and
0.01% bromophenol blue) containing 10 mg/ml
dithiothreitol (Biorad), followed by 15 min in equilibration buffer containing 25 mg/ml iodoacetamide
Latitudinal variation in Geukensia demissa protein expression
639
(Biorad). Strips were placed atop SDS–PAGE gels
and held in place with 0.8% agarose. Gels were run
simultaneously in groups of six or 12 in Tris–glycine
running buffer (25 mM Tris base, 190 mM glycine,
0.1% SDS) using the Biorad Dodeca rig (200 V,
1 h) cooled to 108C. Gels were stained overnight
with colloidal Coomassie blue, and destained with
multiple washes of 18 MV water.
changed significantly between control and HS in
mussels from each collection site.
Gel image analysis
Gel images were recorded at 600 dpi resolution using
an Epson model 700 V transilluminating scanner.
Images were pre-processed using the histogram tool
within the Epson software to maximize contrast, and
were cropped to the gel boundaries using Photoshop.
Sixty gel images were imported into Delta2D image analysis software (DECODON, Greifswald,
Germany), where images were associated with treatment groups, warped (i.e., corresponding spots from
separate gels were associated) and fused. The fusion
image was used to detect spots, and after manual
spot editing, these were transferred back to the original 60 gel images. The relative spot volumes of these
proteins (i.e., the percentage of the total protein
volume on each gel ascribed to each spot) were
used for statistical analysis of changes in protein
abundance in response to each treatment.
Statistical analysis
Before analysis, we removed spots in which mean
relative spot volume was 50.05 in all groups, because
we had difficulty identifying proteins via tandem
mass spectrometry (MS/MS) when protein volume
was below this threshold. Statistical tools within the
multi-experiment viewer module of Delta2D were
used to determine whether PEPs varied in correlation
with latitude among control mussels, and to assess
the impacts of acute HS on protein expression in
gills of G. demissa from each location. A one-way
ANOVA test (P50.01; significance determined by
permutation; standard Bonferroni correction) identified spots changing significantly in abundance between the groups of control (188C-acclimated)
mussels. Hierarchical cluster analysis (HCL), using
spots identified via ANOVA and utilizing average
linking and Pearson’s correlation coefficient, allowed
clustering of individual mussels into discrete groups
based on similarity in PEPs. Clustering of control
mussels by protein expression was further examined
by principal components analysis (PCA). T-tests
(P50.01; significance determined by permutation;
standard Bonferroni correction) were employed to
determine the number of proteins whose abundance
Identification of proteins via tryptic digest and MS/MS
Those protein spots that were identified as having
changed most significantly in response to HS were
excised from gels and destained with 25 mM ammonium bicarbonate in 1:1 (v/v) acetonitrile:water. Gel
pieces were dehydrated by soaking in 100% acetonitrile, and were re-swelled in digestion buffer (25 mM
ammonium bicarbonate) containing 10 ng/mL modified porcine trypsin (Promega). After 4 h incubation
(378C) with shaking, the digest solution was collected for MS analysis.
Tryptic peptides were separated and analyzed
using a high-pressure liquid chromatography
(HPLC)–electrospray ionization ion trap MS/MS
(Agilent 1100 series SL ion trap LC/MS; Santa
Clara, CA, USA). The mobile phase consisted of an
increasing linear gradient of acetonitrile in water,
acidified with formic acid (0.1%); peptides were separated on a nonpolar C8 column (Agilent) before
injection into the trap. Agilent software automatically
detected peptides via MS, and submitted these peptides to MS/MS to allow determination of amino
acid sequences. Peptide mass lists were submitted
to Mascot MS/MS Ions search software (version
3.1; Matrix Science Inc., Boston, MA, USA) for
identification.
Few G. demissa amino acid or nucleotide sequences are available, and so identification of proteins in this study was based on homology, that is, by
matching peptide sequences from G. demissa with
identical sequences from other species. NCBInr and
EST_invertebrates databases were searched for
matches, allowing one missed cleavage by trypsin,
and including cysteinyl carbamidomethylation and
methioninyl oxidation as variable modifications. A
protein was considered identified if Mascot reported
a significant molecular weight search (Mowse) score
(48 for NCBInr; 67 for EST_invertebrates; P50.05),
and included two or more non-overlapping peptides.
For proteins matching a sequence in the EST library,
a subsequent BLAST search was performed to identify the protein by homology.
Results and Discussion
Image analysis of two-dimensional gels
We separated protein samples by two-dimensional
gel electrophoresis (2DGE; Fig. 1) from gill tissue
of six G. demissa (controls; acclimated at 188C)
and six HS G. demissa from each of five locations
(Table 1). Two of the gels, one each from HS
640
P. A. Fields et al.
Fig. 1 Representative 2DGE image of G. demissa gill proteins with spots separated by pI horizontally and molecular mass vertically. Of
the 448 spots detected after applying a low-volume cutoff (0.05), 56 spots that were determined to have changed in abundance in at
least one latitudinal group after HS (t-test, P50.01) are circled. Spots identified by MS/MS are identified by number (Figs. 4–7 and
Supplementary Table S1 for identifications of proteins).
animals from North Carolina (NC) and Virginia
(VA), were of poor quality and were excluded from
analysis. After association of spots across all 58 remaining gels, creation of a composite image, and
removal of all spots with a relative spot volume
50.05, 448 protein spots remained, and were used
for the analyses described below.
Differences in protein expression among Geukensia
demissa acclimated to 188C (controls)
To determine whether differences in protein expression correlated with latitude of origin of the mussel
groups, we examined PEPs in the absence of HS,
using data from control mussels only. Our goal
was to determine whether significant differences in
protein expression remained among mussels from
different locations after acclimation to common conditions for 4 weeks, and if so, whether PEPs varied in
correlation with the latitudes of the sites from which
the mussels were collected. Among the 30 control
mussels, we found 95 proteins that differed significantly in abundance (one-way ANOVA, P50.01;
standard Bonferroni correction) among the five
groups. When expression levels of these proteins
were analyzed by PCA (Fig. 2), mussels from
within collection sites clustered tightly but were
clearly separated from one another. Thus, withingroup variation in protein expression of control
mussels was significantly lower than between-group
variation, indicating that differences in PEPs remained even after acclimation. Importantly, however,
Fig. 2 Principal components analysis of protein abundance in
proteins found to vary among five latitudinal groups of G. demissa.
Ninety-five proteins out of 448 were found to vary significantly
(one-way ANOVA, P50.01) among control (acclimated to 188C)
mussels, and were used for the analysis. Symbols representing
mussels from the same location are outlined and labeled with the
location code (Table 1). Percentages indicate the proportion of
total variation in protein expression explained by each principal
component.
there was no apparent relationship between latitude
of collection site and position of the group clusters
on either the first or second PC, which represented
27.8 and 19.9% of total variation in protein expression, respectively. For example, along PC1 (horizontal axis, Fig. 2) Maine (ME) and VA (Virginia)
groups cluster together to the right and Florida
(FL), NC, and New York (NY) are associated to
the left, although these collection sites are not nearest
neighbors geographically. In contrast, ME and VA
groups are widely separated on PC2 (vertical axis).
Latitudinal variation in Geukensia demissa protein expression
641
The same lack of correlation between protein expression and latitude was found on higher PCs (data not
shown). Similarly, when the 95 significantly varying
proteins were subjected to HCL, mussels clustered
with others from the same collection locality.
However, there was no clear latitudinal relationship
among the clusters (data not shown). From the PCA
and HCL results, we conclude that PEPs vary significantly among the five groups. The persistence of
differences in PEPs after acclimation may be due to
genetic divergence among the mussels from the five
sites or to epigenetic factors; the source of the variability requires further research. It is clear, however,
that protein expression among the groups does not
vary monotonically with latitude.
protein expression among unstressed mussels, exposure to HS might induce changes in protein abundance that varied between northern and southern
G. demissa. Because we were interested in how
protein expression varied between control and HS
in mussels from each collection site, we performed
t-tests (P50.01, standard Bonferroni correction) on
the 448 detected proteins between the two treatments
independently for each location. We found that the
northernmost (ME) population responded more
strongly to HS [Fig. 3; 31 proteins (6.9%) with significantly altered abundance] than did NY or VA
[11 proteins (2.5%)] or the southernmost groups
[NC, five proteins (1.1%); FL, seven proteins (1.6%)].
Of the 65 proteins detected as changing significantly
in abundance in at least one of the groups, there were
56 unique protein spots; the remaining nine changed
significantly in abundance in more than one group.
After tryptic digestion and MS/MS analysis of each
spot, we identified 27 of the 56 proteins (48.2%;
Supplementary Table S1), which could be categorized
into five functional groups: molecular chaperone or
Differences in protein expression after heat stress
We next examined whether protein expression varied
among the mussels from different sites after acute
exposure to a 408C HS. We reasoned that although
we found no latitudinal associated differences in
Fig. 3 Heat maps of gill proteins found to change significantly in abundance after acute HS (408C, 1 h) from G. demissa collected at five
locations across the breadth of the species’ latitudinal range. T-tests (P50.01; standard Bonferroni correction) were performed independently for each location, using expression patterns of 448 proteins detected by 2DGE analysis. Within heat maps, blue color
represents relatively low standardized spot volume and yellow represents relatively high standardized volume. Columns correspond to
individual mussels, with green bars representing controls (n ¼ 6 per site) and red bars representing HS (n ¼ 6 per site, except n ¼ 5 for
NC and VA); rows correspond to expression patterns for individual proteins.
642
folding, cytoskeletal, oxidative stress, cell signaling,
and a single translation initiation factor.
Chaperone proteins
Eleven of the 27 identified proteins were chaperones,
including seven heat-shock protein 70 (HSP70) isoforms (spots 3–9, Fig. 4), two isoforms of small
heat-shock protein 24 (sHSP24; spots 10 and 11),
and two isoforms of 78 kDa glucose-regulated protein [GRP78 (spots 1 and 2), the HSP70 homolog
found in the endoplasmic reticulum (ER)]. One foldase, peptidylprolyl cis–trans isomerase (PPI; spot
22), was also found.
Of the seven HSP70 isoforms, four increased significantly in abundance among ME mussels after HS
and four increased significantly among NY mussels
as well (Fig. 4). In both the NY and ME groups, six
of the seven HSP70 isoforms increased in abundance
more than 1.5-fold (Fig. 4). In contrast, among the
three southern groups, NC had no significant change
in HSP70 levels and VA and FL showed significant
up-regulation of only one HSP70 isoform after HS.
The up-regulation of members of the highly conserved HSP70 family is a ubiquitous indicator of
protein damage due to heat and other environmental
stressors, and HSP70s repeatedly have been found to
P. A. Fields et al.
increase significantly in abundance in mussels and
other mollusks exposed to acute HS (Hofmann and
Somero 1995; Roberts et al. 1997; Tomanek and
Somero 1999; Tomanek and Zuzow 2010). The
408C acute HS to which these mussels were exposed
is close to the 458C upper lethal limit reported for
G. demissa in South Carolina, USA (Jost and
Helmuth 2008), so it is surprising that individuals
from VA, NC, and FL were able to withstand the
treatment with only minimal up-regulation of
HSP70 expression. It is not clear whether their relatively weak response to HS indicates that cellular
damage due to protein denaturation and aggregation
was avoided in these groups, or whether alternate
mechanisms were used to maintain protein structure
and minimize proteotoxic damage.
The two identified sHSP24 isoforms were
up-regulated in ME, NY, and FL groups after HS,
and one of the two (spot 11) significantly increased
in abundance in the VA group as well (Fig. 4).
Again, ME and NY showed the greatest fold-change
in abundance among the five groups. Small
heat-shock proteins play an important role in stabilizing proteins in danger of denaturing during HS
(McHaourab et al. 2009), although they are not
able to refold denatured proteins (Haslbeck et al.
Fig. 4 Average ( SD) relative spot volumes of chaperone proteins from gill tissue of control (open columns) and HS (filled columns)
G. demissa collected from five sites across the species’ latitudinal range (Table 1 for site abbreviations). Proteins were identified by MS/
MS using Mascot search software; spot numbers in graph titles correspond to spots in Fig. 1 (also see Supplementary Table S1).
Numbers above columns indicate the fold-change in average spot volume between control and HS. Asterisks indicate significant
differences (t-test, P50.01, standard Bonferroni correction) between control and HS. Note that y-axis magnitude differs among panels.
643
Latitudinal variation in Geukensia demissa protein expression
Fig. 5 Average ( SD) abundances of cytoskeletal proteins from gill tissue of control (open columns) and HS (filled columns)
G. demissa. Data are presented as described in Fig. 4.
2005). In light of the changes in actin expression in
G. demissa after HS (described below), it is interesting to note that sHSPs also have been found to interact with the actin cytoskeleton during various
stresses, including HS and oxidative stress (Arrigo
1998; Dalle-Donne et al. 2001; Stromer et al. 2003).
It appears that sHsps have the ability both to inhibit
the polymerization of filamentous actin during cytotoxic stress (Wieske et al. 2001), and also limit disruption and aggregation of actin filaments (Mounier
and Arrigo 2002).
The two ER chaperone proteins identified, isoforms of GRP78, were relatively unchanged in abundance after HS for most groups. One GRP78 was
significantly down-regulated in both FL and ME
groups (spot 02, Fig. 4), while the other (spot 01)
was significantly up-regulated in the ME group only.
GRP78 is up-regulated during ER stress (Rao et al.
2002), and is involved in Ca2þ homeostasis and apoptosis as well as in performing chaperone functions.
Given the central role of GRP78 in mediation of the
unfolded protein response (UPR) in the ER (Lai
et al. 2010), the down-regulation or lack of change
in abundance in these isoforms in all but one group
of G. demissa is unexpected. The up-regulation of
spot 01 in the ME group, in contrast, suggests a
lower threshold for UPR in the ER of gills in these
mussels.
The final chaperone/foldase protein we identified
from G. demissa gill, PPI, is also found in the ER and
catalyzes the rate-limiting step in protein folding,
interconversion of cis and trans isomers of proline
(Budiman et al. 2011). This chaperone was
up-regulated nearly two-fold in both the VA and
ME groups, suggesting increased protein synthesis
or refolding in the ER.
In summary, the ME mussels appeared to have the
most significant up-regulation of both cytosolic and
ER chaperone and folding proteins after HS, with
nine of 12 proteins changing significantly in abundance. In comparison, six of 12 increased significantly in abundance in the neighboring NY group.
In most cases, isoforms from these two northernmost
groups also experienced the greatest fold-change in
abundance. Taken together, these data suggest that
the ME and NY groups more readily induce a significant stress response after HS compared to the
three southern groups.
Cytoskeletal proteins
Ten proteins associated with the cytoskeleton were
identified, and abundance changed significantly
only within the ME group in all but two (Fig. 5).
One b-tubulin (spot 12) isoform and one a-tubulin
(spot 13) isoform were found; the former was expressed at 75% of its original abundance after HS
in the ME mussels; the latter increased in abundance
more than three-fold. In addition, six actin isoforms
altered abundance significantly, but only in the ME
group. Three isoforms (14, 15, and 16) decreased in
644
P. A. Fields et al.
Fig. 6 Average ( SD) abundances of two isoforms of glutaredoxin from gill tissue of control (open columns) and HS (filled columns)
G. demissa. Data are presented as described in Fig. 4.
abundance by 450% while three others (17, 18, and
19) increased in abundance two-fold or more.
Interestingly, the down-regulated isoforms appear at
a slightly higher mass and more acidic pI relative to
the up-regulated ones (Fig. 1). A final actin isoform
(spot 20) was chosen for identification because of its
proximity to other significantly changing spots on
the gels, but there was no significant change in abundance in any group for this isoform, despite an overall pattern of increasing expression after HS.
The final cytoskeleton-associated protein identified
was F-actin capping protein subunit a (spot 21).
Capping protein (CP) binds to fast-growing ends
of actin filaments, stabilizing the structure and inhibiting addition or loss of actin monomers (Wear
and Cooper 2004). F-actin capping protein was significantly down-regulated after HS in the NY group.
The changes in abundance of actin isoforms in the
ME mussels and CP in the NY mussels suggests considerable modification of the actin cytoskeleton in
response to HS in these groups, and in ME most
significantly. Changes in abundance of cytoskeletal
protein, including substantial changes in actin expression, have been found in other mussels exposed
to HS (Mytilus trossulus and M. galloprovincialis;
Tomanek and Zuzow 2010), and there is strong evidence that actin filaments are disrupted both by heat
(Glass et al. 1985; Welch and Suhan 1985) and redox
stress (McDonagh and Sheehan 2008). Furthermore,
it is clear that sHSPs modulate assembly of actins
(Liang and MacRae 1997), and that during stress
sHSPs inhibit polymerization of actin and modify
the stress fibers and focal adhesions through which
cells interact with the extracellular matrix (Schneider
et al. 1998). The PEPs of actin and sHSPs in the ME
mussels, and CP and sHSPs in the NY group, support the model proposed by Tomanek and Zuzow
(2010), in which cytoskeletal elements, including
actin-based microfilaments, are destabilized by HS,
and cells respond to the disruption both by modulating abundance of actin isoforms and by inducing
synthesis of sHSPs. Notably, however, in G. demissa,
this process is only observed in the two northernmost groups, and most significantly among ME
mussels.
Oxidative stress proteins
Two isoforms of the oxidative stress protein glutaredoxin (GRX) were identified via MS/MS (spots 23
and 24), and appear to differ slightly in MW based
on their location on the 2D gels (Fig. 1). The close
apposition of the spots is suggestive of
post-translational modification of GRX affecting
MW but not pI, although the limited sequence coverage of the molecules obtained by MS/MS
(Supplementary Table S1) is not sufficient to confirm this. The lower MW isoform (spot 23) was
found to increase significantly in abundance after
HS, but only in VA and ME mussels (Fig. 6); the
seven-fold increase in this GRX isoform in the ME
mussels was the largest change in abundance found
for any protein after HS. In contrast, the larger GRX
isoform (spot 24) decreased significantly after HS
only in the ME group.
GRXs possess active-site thiol groups that are reduced by glutathione, which is then re-reduced by
glutathione reductase (Holmgren 1989). The thiols
allow GRXs to act as disulfide reductases or reductases of protein-mixed disulfides created during oxidative stress (Grant 2001). In this context, it is
interesting to note that one of the main targets for
glutathionylation during oxidative stress is the actin
cytoskeleton (McDonagh et al. 2005), forming
protein-mixed disulfides. Oxidative damage is induced by multiple types of cellular stress, including
HS (Kültz 2005), and thus modification of expression of GRX, co-occurring with changes in abundance of actin isoforms and sHSP24s, may also
play a role in maintaining cytoskeletal structure in
response to heat-induced oxidative stress. Again, it is
noteworthy that this response is most pronounced in
the ME mussels.
645
Latitudinal variation in Geukensia demissa protein expression
Fig. 7 Average ( SD) abundances of two isoforms of the regulatory protein 14-3-3, as well as translation initiation factor 5A, from gill
tissue of control (open columns) and HS (filled columns) G. demissa. Data are presented as described in Fig. 4.
Cell signaling and transcription
The cell signaling protein 14-3-3 was detected in two
spots (25 and 26), which occurred at significantly
different positions on the gels (Fig. 1). The protein
with a higher MW (spot 25) is closer to the expected
14-3-3 MW of 29 kDa, and shows a greater than
three-fold up-regulation in the ME mussels after
HS (Fig. 7). The second 14-3-3 spot detected (26)
varies in expression depending on the group tested,
decreasing significantly in abundance among NY
mussels, but increasing significantly among VA mussels. However, because this spot occurs at 15 kDa
on the gels, it may represent a proteolytic artifact
whose variation does not reflect a cellular response
to HS.
The 14-3-3 proteins are highly conserved in eukaryotes, and are able to bind a wide variety of signaling proteins including kinases, phosphatases,
transmembrane receptors, and transcription factors
(Fu et al. 2000). Because of their ubiquity, 14-3-3
proteins have been implicated in the control of a
variety of cellular functions, from cell cycle to metabolism and apoptosis (Bridges and Moorhead
2005). Because of the diversity of roles ascribed to
14-3-3 proteins, the cellular role of the isoform
up-regulated in the ME mussels cannot be determined. This change in 14-3-3 expression in the ME
mussels compared to the other groups, however,
provides further evidence that cellular processes are
most heavily impacted by HS in the northernmost
group.
The final protein identified is eukaryotic transcription initiation factor 5A (eIF5A) (spot 27, Fig. 7).
eIF5A has a central role in transcription initiation,
interacting with the 40S initiation complex and hydrolyzing bound GTP to allow release of eIF2 and
subsequent binding of the 60S ribosomal subunit
(Das et al. 2001). In G. demissa, the abundance of
eIF5A decreases significantly after HS only in the NC
group, and only to a level about 75% that of controls. The apparent MW of the gel spot, 25 kDa,
is substantially lower than the expected MW of
46–49 kDa (representing the mass of the protein in
yeast and mammals, respectively; Das et al. 2001),
suggesting again that this spot may be a cleavage
product of a larger protein. Based on these observations, we conclude that eIF5A does not play an
important role in response to HS, despite its identification here.
Conclusions
The proteomic analysis we describe here suggests
strongly that G. demissa from ME, and to a lesser
extent NY, are more strongly affected by HS than are
conspecifics collected farther south in the species’
extensive latitudinal range. In this context, it is noteworthy that Cape Cod represents a significant biogeographic barrier and the Gulf of Maine is
thermally distinct from the Atlantic Ocean south of
the Cape (Table 1) (Vermeij 1978; Bertness et al.
1999). The ME mussels collected north of this discontinuity showed substantially greater up-regulation
646
of chaperone and cytoskeletal proteins, indicating a
greater level of protein denaturation and structural
damage. We also found evidence for greater sensitivity to heat-induced oxidative stress and a suggestion
of more substantial modification of cellular processes
after HS.
Taken together, these data indicate that there are
significant physiological differences in G. demissa
based on geographic location, which are not removed through acclimation to common conditions.
Our results are consistent with localized adaptation
to warmer conditions in the southern portion of the
range, and cooler conditions in the northern part,
especially north of Cape Cod. It remains to be determined whether the source of these differences is
genetic or epigenetic, or something else altogether.
Acknowledgments
We thank L. Tomanek for the invitation to present
this work in the Comparative Environmental
Proteomics symposium held at SICB 2012. We
would also like to express our gratitude to Dr Ken
Hess for his help in mass spectrometry.
Funding
This work was supported by the National Science
Foundation (grant IOS 0920103) as well as a grant
to Franklin & Marshall College from the Howard
Hughes Medical Institute.
Supplementary material
Supplementary material is available at ICB online.
References
Arrigo AP. 1998. Small stress proteins: chaperones that act as
regulators of intracellular redox state and programmed cell
death. Biol Chem 379:19–26.
Bertness MD. 1984. Ribbed mussels and Spartina alterniflora
production in a New England salt marsh. Ecology
65:1794–807.
Bertness MD, Leonard GH, Levine JM, Bruno JF. 1999.
Climate-driven interactions among rocky intertidal organisms caught between a rock and a hot place. Oecologia
120:446–50.
Bertness MD, Grosholz E. 1985. Population dynamics of the
ribbed mussel, Geukensia demissa: the costs and benefits of
an aggregated distribution. Oecologia 67:192–204.
Bridges D, Moorhead GBG. 2005. 14-3-3 proteins: A number
of functions for a numbered protein. Sci STKE 2005:re10.
Budiman C, Koga Y, Takano K, Kanaya S. 2011.
FK506-binding protein 22 from a psychrophilic bacterium,
a cold shock-inducible peptidyl prolyl isomerase with the
ability to assist in protein folding. Int J Mol Sci 12:5261–84.
Dalle-Donne I, Rossi R, Milzani A, Di Simplicio P,
Colombo R. 2001. The actin cytoskeleton response to
P. A. Fields et al.
oxidants: from small heat shock protein phosphorylation
to changes in the redox state of actin itself. Free Radic
Biol Med 12:1624–32.
Das S, Ghosh R, Maitra U. 2001. Eukaryotic translation initiation factor 5 functions as a GTPase-activating protein. J
Biol Chem 276:6720–6.
Fu H, Subramanian RR, Masters SC. 2000. 14-3-3 proteins:
structure, function, and regulation. Annu Rev Pharmacol
Toxicol 40:617–47.
Glass JR, DeWitt RG, Cress AE. 1985. Rapid loss of stress
fibers in Chinese hamster ovary cells after hyperthermia.
Cancer Res 45:258–62.
Gliman SE, Wethey DS, Helmuth B. 2006. Variation in the
sensitivity of organismal body temperature to climate
change over local and geographic scales. Proc Natl Acad
Sci USA 103:9560–5.
Gorg A. 2004. 2-D electrophoresis: principles and methods.
Little Chalfont, UK: GE Healthcare.
Gosner KL. 1978. A field guide to the Atlantic seashore. New
York, NY: Houghton Mifflin Co.
Grant CM. 2001. Role of the glutathione/glutaredoxin and
thioredoxin systems in yeast growth and response to
stress conditions. Mol Microbiol 39:533–41.
Haslbeck M, Franzmann T, Weinfurtner D, Buchner J. 2005.
Some like it hot: the structure and function of small
heat-shock proteins. Nat Struct Mol Biol 12:842–6.
Helmuth B, Broitman BR, Yamane L, Gilman SE, Mach K,
Mislan KAS, Denny MW. 2009. Organismal climatology:
Analyzing environmental variability at scales relevant to
physiological stress. J Exp Biol 213:995–1003.
Helmuth B, Harley CDG, Halpin PM, O’Donnell M,
Hofmann GE, Blanchette CA. 2002. Climate change and
latitudinal patterns of intertidal thermal stress. Science
298:1015–7.
Hofmann GE, Somero GN. 1995. Evidence for protein
damage at environmental temperatures: Seasonal changes
in levels of ubiquitin conjugates and HSP70 in the intertidal mussel Mytilus trossulus. J Exp Biol 7:1509–18.
Holmgren A. 1989. Thioredoxin and glutaredoxin systems. J
Biol Chem 264:13963–6.
Hutchins LW. 1947. The bases for temperature zonation in
geographical distribution. Ecol Monogr 17:325–35.
Jones SJ, Lima FP, Wethey DS. 2010. Rising environmental
temperatures and biogeography: poleward range contraction of the blue mussel, Mytilus edulis L., in the western
Atlantic. J Biogeogr 37:2243–59.
Jost J, Helmuth B. 2008. Morphological and ecological determinants of body temperature of Geukensia demissa, the
Atlantic Ribbed Mussel, and their effects on mussel mortality. Biol Bull 213:141–51.
Kanwisher JW. 1955. Freezing in intertidal animals. Biol Bull
109:56–63.
Kültz D. 2005. Molecular and evolutionary basis of the cellular stress response. Annu Rev Physiol 67:225–57.
Lai CW, Aronson DE, Snapp EL. 2010. BiP availability
distinguishes states of homeostasis and stress in the
endoplasmic reticulum of living cells. Mol Biol Cell
21:1909–21.
Lent CM. 1968. Air-gaping by the ribbed mussel, Modiolus
demissus (Dillwyn): effects and adaptive significance. Biol
Bull 134:60–73.
Latitudinal variation in Geukensia demissa protein expression
647
Lent CM. 1969. Adaptations of the ribbed mussel, Modiolus
demissus (Dillwyn), to the intertidal habitat. Am Zool
9:283–92.
Liang P, MacRae TH. 1997. Molecular chaperones and the
cytoskeleton. J Cell Sci 110:1431–40.
McDonagh B, Sheehan D. 2008. Effects of oxidative stress on
protein thiols and disulphides in Mytilus edulis revealed by
proteomics: actin and protein disulphide isomerase are
redox targets. Mar Environ Res 66:193–5.
McDonagh B, Tyther R, Sheehan D. 2005. Carbonylation and
glutathionylation of proteins in the blue mussel Mytilus
edulis detected by proteomic analysis and Western blotting:
actin as a target for oxidative stress. Aquat Toxicol
73:315–26.
McHaourab HS, Godar JA, Stewart PL. 2009. Structure and
mechanism of protein stability sensors: chaperone activity
of small heat shock proteins. Biochemistry 48:3828–37.
Mounier N, Arrigo AP. 2002. Actin cytoskeleton and small
heat shock proteins: how do they interact? Cell Stress
Chaperones 2:167–76.
Osovitz CJ, Hofmann GE. 2007. Marine macrophysiology:
Studying physiological variation across large spatial scales
in marine systems. Comp Biochem Physiol A 147:821–7.
Rao RV, Peel A, Logvinova A, del Rio G, Hermel E, Yokota T,
Goldsmith PC, Ellerby LM, Ellerby HM, Bredesen DE.
2002. Coupling endoplasmic reticulum stress to the cell
death program: role of the ER chaperone GRP78. FEBS
Lett 514:122–8.
Roberts D, Hofmann GE, Somero GN. 1997. Heat-shock protein expression in Mytilus californianus: acclimatization
(seasonal and tidal-height comparisons) and acclimation
effects. Biol Bull 192:309–20.
Schneider GB, Hamano H, Cooper LF. 1998. In vivo evaluation of hsp27 as an inhibitor of actin polymerization:
Hsp27 limits actin stress fiber and focal adhesion formation
after heat shock. J Cell Physiol 177:575–84.
Somero GN. 2005. Linking biogeography to physiology:
evolutionary and acclimatory adjustments of thermal
limits. Front Zool 2:1–9.
Sorte CJB, Hofmann GE. 2005. Thermotolerance and
heat-shock protein expression in Northeastern Pacific
Nucella species with different biogeographic ranges. Mar
Biol 146:985–93.
Stillman J. 2003. Acclimation capacity underlies susceptibility
to climate change. Science 301:65.
Stromer T, Ehrnsperger M, Gaestel M, Buchner J. 2003.
Analysis of the interaction of small heat shock proteins
with unfolding proteins. J Biol Chem 278:18015–21.
Tomanek L. 2011. Environmental proteomics: Changes in the
proteome of marine organisms in response to environmental stress, pollutants, infection, symbiosis, and development.
Annu Rev Mar Sci 3:373–99.
Tomanek L, Somero GN. 1999. Evolutionary and
acclimation-induced variation in the heat-shock responses
of congeneric marine snails (genus Tegula) from different
thermal habitats: implications for limits of thermotolerance
and biogeography. J Exp Biol 202:2925–36.
Tomanek L, Zuzow MJ. 2010. The proteomic response of the
mussel congeners Mytilus galloprovincialis and M. trossulus
to acute heat stress: implications for thermal tolerance
limits and metabolic costs of thermal stress. J Exp Biol
213:3559–74.
Vermeij GJ. 1978. Biogeography and adaptation: patterns of
marine life. Cambridge, MA: Harvard University Press.
Wear MA, Cooper JA. 2004. Capping protein: new insights
into mechanism and regulation. Trends Biochem Sci
29:418–28.
Welch WJ, Suhan JP. 1985. Morphological study of the mammalian stress response: characterization of changes in cytoplasmic organelles, cytoskeleton, and nucleoli, and
appearance of intranuclear actin filaments in rat fibroblasts
after heat-shock treatment. J Cell Biol 101:1198–211.
Wells HW. 1961. The fauna of oyster beds with special reference to the salinity factor. Ecol Monogr 31:329–66.
Wethey DS. 1983. Geographic limits and local zonation: the
barnacles Semibalanus (Balanus) and Chthalamus in New
England. Biol Bull 165:330–41.
Wieske M, Benndorf R, Behlke J, Dölling R, Grelle G,
Bielka H, Lutsch G. 2001. Defined sequence segments of
the small heat shock proteins HSP25 and alphaB-crystallin
inhibit actin polymerization. Eur J Biochem 268:2083–90.