Effects of High Hydrostatic Pressure on Coastal Bacterial Community Abundance and Diversity

Abstract

Hydrostatic pressure is an important parameter influencing the distribution of microbial life in the ocean. In this study, the response of marine bacterial populations from surface waters to pressures representative of those under deep-sea conditions was examined. Southern California coastal seawater collected 5 m below the sea surface was incubated in microcosms, using a range of temperatures (16 to 3°C) and hydrostatic pressure conditions (0.1 to 80 MPa). Cell abundance decreased in response to pressure, while diversity increased. The morphology of the community also changed with pressurization to a predominant morphotype of small cocci. The pressure-induced community changes included an increase in the relative abundance of Alphaproteobacteria, Gammaproteobacteria, Actinobacteria, and Flavobacteria largely at the expense of Epsilonproteobacteria. Culturable high-pressure-surviving bacteria were obtained and found to be phylogenetically similar to isolates from cold and/or deep-sea environments. These results provide novel insights into the response of surface water bacteria to changes in hydrostatic pressure.

INTRODUCTION

Hydrostatic pressure constitutes an important but understudied parameter influencing life on Earth. High-pressure environments include the deep sea and the deep subsurface, wherein exists a plethora of life encompassing a major fraction of the biosphere (1, 2). High hydrostatic pressure inhibits cellular processes and the formation of macromolecular structures that result in a positive volume change. In mesophilic microorganisms, these processes include motility, substrate transport, cell division, DNA replication, transcription, translation, certain enzymatic reactions, and changes in membrane liquid crystalline states (3–10). In the case of the mesophile Escherichia coli, exposure to high pressure results in the inhibition of cell division, DNA replication, and protein synthesis and the paradoxical induction of both cold shock and heat shock proteins (7). Even though there is considerable overlap between the adaptation to low temperature and high pressure, there are also major differences, as revealed by analyses of cold-sensitive and pressure-sensitive mutants derived from high-pressure-adapted (piezophilic) Photobacterium profundum strain SS9 (11). Many cold-sensitive mutants are not pressure resistant and vice versa, underscoring the unique nature of adaptation to high pressure (11).

The dynamic nature of the marine environment can hinder efforts to study the diversity, abundance, and distribution of microbes across zones distinguishable by their physical and chemical properties. Ocean mixing occurs over a wide range of vertical and horizontal scales (12, 13). Sinking organic detritus from the surface of the ocean, known as marine snow, also contributes to the mixing and introduces allochthonous microbes and nutrients to the deep ocean. Examples that reinforce the view of the ocean as a dynamic system include the recovery of cyanobacterial communities from the deep sea and the recovery of thermophilic bacteria from cold Arctic sediments (14–16).

The role of hydrostatic pressure in shaping the activity and distribution of microbial life in the ocean has been acknowledged in studies dating back to 1884 (17). ZoBell and Johnson (18) and ZoBell and Oppenheimer (19) reported that high hydrostatic pressure (20 to 60 MPa) inhibited the growth of surface marine bacteria at 30°C, while low temperature exacerbated the inhibitory effects of pressure. Another study 25 years later investigated the ability of sewage bacteria to survive deep-sea conditions and concluded that certain microbes found in raw sewage, including E. coli and Enterococcus faecalis, were able to tolerate long-term exposure to pressures up to 50 MPa at 4°C (20). More recently, two independent groups addressed the effect of pressure and low temperature on surface microbial communities, focusing on either selected species or microbial assemblages associated with marine snow (21–23). Grossart and Gust studied the response of five shallow-water strains to a pressurized microcosm simulating the pressure at a 4,000-m depth and reported a strong pressure response, reflected by changes in the relative abundance of the examined strains (21). Tamburini et al. reported that pressure affected both the activity and the relative abundance of the prokaryotes associated with sinking organic matter, detecting a pressure-dependent increase in the relative abundance of Gammaproteobacteria over time (22, 23).

In this study, additional dimensions of elevated pressure effects on shallow-water bacterioplankton were investigated over exposures up to 80 MPa and periods up to 1 month. Samples were collected from a coastal environment in Southern California, and the microbial community changes as a function of pressure were evaluated using 16S rRNA gene analysis. In addition, we were able to isolate and characterize high-pressure-tolerant bacteria.

MATERIALS AND METHODS

Sample collection and incubation conditions.

Seawater samples were collected from the end of the pier at the Scripps Institution of Oceanography (32°52′N, 117°15′W) at an approximately 5-m depth below sea level. The sea temperature (at 5 m below sea level), salinity, pressure, and chlorophyll II concentrations were obtained at the time of collection through the Southern California Coastal Ocean Observing System (http://www.sccoos.org/). (See Table S1 in the supplemental material for sample collection dates and the reported seawater temperature for each experiment.) Seawater samples from each collection date were pooled and stored for 14 days at 16°C in the dark until further processing.

All seawater samples (150 to 200 ml) were filtered through a 0.8-μm-pore-size membrane filter (Pall Corporation, San Diego, CA) to remove large particles and predators (24, 25). Replicates of the filtered seawater were placed in sterilized Kapak bags (Kapak Corporation, Minneapolis, MN) and incubated in the dark at the corresponding temperature (16°C or 3°C) and hydrostatic pressure (0.1, 10, 30, 60, and 80 MPa), while stainless steel pressure vessels were used for both low- and high-pressure treatments. Seawater was also used as the inoculum (10%) to set up enrichment cultures in 75%-strength marine broth 2216 medium (BD Difco) in 5-ml polyethylene transfer pipette bulbs (Samco Scientific, San Fernando, CA), and the cultures were incubated in the dark at 16°C at 0.1, 30, and 60 MPa.

Following depressurization, aliquots from each treatment were plated on marine agar 2216 (BD Difco) to determine the number of bacterial CFU ml⁻¹ of sample. The plates were incubated at 16°C in the dark at atmospheric pressure. Colonies from high-pressure-surviving bacteria were selected for further characterization. The growth characteristics of the high-pressure-surviving isolates were determined by growing the selected isolates in 5-ml polyethylene transfer pipette bulbs with 75%-strength marine broth 2216 at 16°C in the dark at 0.1, 20, 40, and 60 MPa. The piezophile Photobacterium profundum SS9 was used as a positive control and was grown in 75%-strength marine broth 2216 at 16°C in the dark. The nonpiezophile Escherichia coli was used as a negative control and was grown in Luria-Bertani (LB) medium at 37°C in the dark.

DNA isolation.

Seawater samples were filtered through a 0.1-μm-pore-size membrane filter (Pall Corporation, San Diego, CA) for DNA or RNA extraction (26, 27). Filters for DNA extraction were submerged in lysis buffer (20 mM EDTA, 0.4 M NaCl, 0.75 M sucrose, 50 mM Tris-HCl, 2 mg ml⁻¹ lysozyme) for 1 h, followed by proteinase K (200 μg ml⁻¹) and 1% SDS for 2 h. The DNA was extracted with phenol-chloroform and precipitated with sodium acetate-ethanol. The DNA pellet was resuspended in TE (Tris-EDTA) buffer.

16S rRNA gene clone library construction and sequencing.

DNA (6 ng) isolated from seawater samples incubated at either 0.1 MPa or 60 MPa (3°C, in the dark) for 14 days was used as the template for the amplification of the bacterial 16S rRNA gene using the universal primers 27F and 1492R (28). The PCR was performed with Platinum Taq High-Fidelity DNA polymerase (Life Technologies). The cycling program was performed as follows: 5 min at 95°C, followed by 35 cycles of 30 s at 94°C, 2 min at 52°C, and 2 min at 72°C and extension at 72°C for 10 min. Following amplification, A tails were added and the PCR products were purified using a QIAquick gel extraction kit (Qiagen) according to the manufacturer's instructions. The purified PCR products were ligated into the pGEM-T Easy cloning vector (Promega) and transformed into E. coli JM109 cells. Selected clones were submitted for direct sequencing (Beckman Coulter Genomics). A total of 115 and 112 16S rRNA clones were sequenced for the samples incubated at 0.1 MPa and 60 MPa, respectively. The sequences were aligned using the NAST alignment tool and checked for chimeras using the Bellerophon program (v.3) (29). The Library Compare tool was used to compare the two microbial communities using a naive Bayesian classifier (30). The program Metastats was used to statistically test the significant differences at the level of order grouping between the two libraries (31).

16S rRNA gene sequencing was also performed on isolated colonies. In this case, a single colony of a high-pressure-surviving isolate was placed in 100 μl of Milli-Q water, resuspended by vortexing, and heated at 96°C for 10 min. The supernatant (2 μl) was used as the template for the amplification of the16S rRNA gene using the 27F and 1492R primers. The PCR products were submitted for direct sequencing (SeqXcel, San Diego, CA). The sequences were aligned using the NAST alignment tool and checked for chimeras using Bellerophon (v.3) (29). The Phylemon (v.2.0) suite of molecular tools was utilized for sequence trimming and construction of a phylogenetic tree; sequences were trimmed using the TrimAl (v.1.3) program and exported for bootstrap analysis using the maximum likelihood methods implemented in the PhyML package (32–34). The nearest-neighbor interchange method implemented in PhyML was used to construct a phylogenetic tree, while the FigTree viewer was used for tree visualization (http://tree.bio.ed.ac.uk/software/figtree/). 16S rRNA gene sequences for the phylogenetic tree analysis were selected from a comprehensive list of piezotolerant and piezophilic isolates published by Eloe et al., and Marinitoga piezophila KA3 was selected as the outgroup (35). Sequence similarity analysis was performed with the EMBOSS Needle pairwise sequence alignment tool (36).

T-RFLP.

16S rRNA gene sequences were amplified with primer 27F labeled at the 5′ end with 6-carboxyfluorescein (Life Technologies) and primer 1492R with the cycling parameters described above. Reactions with three independent 50-μl reaction mixtures were performed for each sample using 5 to 7 ng of template DNA per reaction mixture. Purified PCR products were digested for 2.5 h (total volume, 50 μl) using the HaeIII restriction enzyme. Digests were precipitated in ethanol with sodium acetate and resuspended in 10 μl of Milli-Q water. Fragment analysis was performed with a 3730xl genetic analyzer (Applied Biosystems, Foster City, CA) and a GeneScan 500 LIZ size standard by Genewiz, South Plainfield, NJ.

Terminal restriction fragment (TRF) length and abundance were determined with GeneMarker (v.1.7) software (Softgenetics, State College, PA) using the minimal peak detection threshold of 50 fluorescent units. Terminal restriction fragment length polymorphism (T-RFLP) data were denoised, binned, and normalized in R as described by Abdo et al. using a cutoff value of two times the standard deviation (37). Since the TRFs were generated with the same 16S rRNA primers used for the 16S rRNA clone libraries, we were able to putatively identify their taxonomy by comparing their fragment size to the sizes of the in silico-digested 16S rRNA sequences.

Microscopy.

Cell counts were determined microscopically. Samples were stained using 4′,6′-diamidino-2-phenylindole (DAPI) nucleic acid stain (Vector Laboratories, Inc., Burlingame, CA) and viewed at ×1,000 magnification on an Olympus BX51 fluorescence microscope (Olympus). Images were processed with ImageJ software (http://rsbweb.nih.gov/ij/).

Statistical analysis.

The Shannon-Weaver diversity index (H′) was computed using the diversity result function in the R package BiodiversityR on the basis of the relative abundance of TRFs in each sample or the 16S rRNA library clone sequences (38). The Bray-Curtis similarity index (including the presence/absence and relative abundance of TRFs) was employed to cluster the communities, using the hclust function in the R package vegan, on the basis of the average linkage method (39). The Bray-Curtis similarity index was also used for nonmetric multidimensional scaling (nMDS) analysis, with the environmental factors fitted using a linear regression using the vegan package. Permutational multivariate analysis of variance was performed using the Adonis function in the vegan package to evaluate the environmental and experimental parameters affecting the community variation observed in the nMDS results.

Graphs were plotted and analyzed using Prism (v.6) software (GraphPad, La Jolla, CA), and statistical analysis was performed by nonparametric 2-tailed Mann-Whitney U test.

Nucleotide sequence accession numbers.

The 16S rRNA sequences of the libraries clone obtained at pressures of 0.1 MPa and 60 MPa were deposited in GenBank under accession numbers KJ002083 to KJ002190 and KJ002191 to KJ002298, respectively. The 16S rRNA sequences of the high-pressure-surviving isolates were deposited in GenBank under accession numbers JX293349 to JX293353.

RESULTS

Microbial abundance and morphology.

In order to begin addressing the effect of elevated pressure and low temperature on the abundance of a coastal microbial community, seawater samples were incubated for different lengths of time at 0.1 and 60 MPa at 3°C in the dark in order to simulate the physical conditions encountered at a depth of ∼6,000 m. The cell numbers in control samples (pretreatment, filtered [pore size, 0.8 μm] seawater) averaged 5 × 10⁵ cells ml⁻¹. The cell numbers remained relatively unchanged for both the samples held at atmospheric pressure and the samples held at elevated pressure for up to 4 days (Fig. 1A). However, a more than ∼50% decrease in cell numbers was observed at the 8th day of incubation for the 60-MPa incubations compared to the cell numbers in the pretreatment samples. A more than 2-fold increase in cell numbers to ∼1.2 × 10⁶ cells ml⁻¹ was also observed from 14 to 56 days of incubation for samples incubated at 0.1 MPa. The high-pressure sample's cell abundance stabilized from 14 to 56 days to approximately 2.5 × 10⁵ cells ml⁻¹. These results indicate that the 60-MPa pressurization of an unamended seawater microcosm produced a net difference in cell abundance of ∼400% compared to that of an identical microcosm incubated at 0.1 MPa.

FIG 1.

Bacterial cell abundance at high pressure. Cell counts were determined microscopically as a function of incubation time at 3°C (A), pressure (B), sampling season (C), and temperature for 14 days (D). The control (Cntrl) was filtered (pore size, 0.8 μm) untreated seawater at day 0. Bar histograms indicate the mean value of at least three replicates, and the error bars indicate the standard deviations of the means.

In order to assess the impact of a wider range of pressures, 2-week microcosms were established under conditions identical to those described above but at 0.1, 10, 30, 60, and 80 MPa (Fig. 1B). A 100-fold increase in pressure (10 MPa), corresponding to that at a water column depth of 1,000 m, caused a considerable decrease in cell abundance. Cell numbers progressively decreased with increasing pressure until at 80 MPa only about a third of the pretreatment cell numbers persisted. The observed decrease in cell numbers was more gradual, with higher cell abundances being found at 10 and 30 MPa for samples collected in January 2010. Even though the sample collection date did not affect the relative abundance trend observed between samples incubated at atmospheric pressure and 60 MPa, higher cell numbers were observed at high pressure for samples collected in January (Fig. 1C). Incubation temperature also affected microbial abundance (Fig. 1D). At atmospheric pressure, lower temperatures, such as 8°C and 3°C, caused modest decreases in cell abundance, but these impacts were far less pronounced than the impact of a 60-MPa treatment at all of the temperatures tested.

DAPI-stained fluorescent images of the microcosm-associated microorganisms present in seawater incubated at 3°C for 14 days at 0.1 MPa, 30 MPa, or 60 MPa are presented in Fig. 2A. Significant changes in the morphology of the microbial community were observed at high hydrostatic pressure. Samples incubated at atmospheric pressure displayed a wide variety of shapes and sizes, while the morphology of those incubated at 30 MPa was less diverse, with the majority of the microbial cells being coccoid. At 60 MPa, the only morphology observed was small cocci, with cell diameters being between 0.6 and 0.2 μm.

FIG 2.

DAPI-stained micrographs of seawater and seawater enrichments incubated at 0.1 MPa, 30 MPa, or 60 MPa for 14 days in the dark. (A) Seawater was incubated at 3°C; (B) seawater was inoculated (10%) in marine broth (MB) and incubated at 16°C.

Enrichment cultures were also prepared in marine broth and incubated at 0.1 MPa, 30 MPa, and 60 MPa at 16°C for 14 days in the dark (Fig. 2B). Under these conditions cell numbers increased dramatically at both 0.1 and 30 MPa. At 0.1 MPa, there was once again a plethora of cell sizes and shapes, but at 30 MPa, the majority of the cells were filamentous, thus exhibiting the well-documented supraoptimal pressure stress response exhibited by many bacteria under conditions in which cell division is inhibited more than growth (5, 19, 40–43). When the enrichments were incubated at 60 MPa, the prominent morphology was small cocci, as was observed in the unamended samples (Fig. 2A). No cell filamentation was ever observed under atmospheric pressure conditions even at low temperatures (data not shown). Live/dead staining confirmed the presence of viable cells in samples incubated at 3°C for 14 days at 0.1 MPa or 60 MPa in seawater (see Fig. S1 in the supplemental material). Scanning electron microscopy indicated the presence of diverse cell morphologies at 0.1 MPa, with cell diameters being between 0.3 to 0.8 μm, while at 60 MPa the majority of the cells were cocci, with cell diameters being reduced to 0.2 to 0.3 μm (see Fig. S1 in the supplemental material).

Community structure.

Changes in the bacterial community structure as a function of pressure, temperature, incubation period, and season were examined by 16S rRNA gene terminal restriction fragment polymorphism (T-RFLP). Community similarity dendrograms were prepared using the T-RFLP fragment clusters (absence/presence and relative abundance) for the tested microcosms (Fig. 3). Samples incubated for 2 days at atmospheric pressure and high pressure clustered together, while samples incubated for 8, 14, or 28 days clustered according to pressure (Fig. 3A). Analysis of the effect of incubation time on community composition indicated the presence of three clusters (Fig. 3A). The T-RFLP profiles for the samples incubated at high pressure for 2 days clustered with those for the samples incubated at high pressure for 8, 14, and 28 days. The control samples formed a separate cluster with a high similarity to the cluster incubated at 60 MPa for 2 days, while the samples incubated at atmospheric pressure (for 8, 14, and 28 days) clustered separately, showing an increased heterogeneity in bacterial composition between the different incubation dates. The overall clustering pattern reflected changes in community diversity; samples with higher diversity (samples incubated at 60 MPa and control samples) clustered together (see Fig. S2B in the supplemental material).

FIG 3.

Dendrograms illustrating bacterial community similarity. Bray-Curtis similarity distance matrices were used for the construction of dendrograms, obtained by the unweighted-pair group method using average linkages, representing relative abundance data from T-RFLP analysis of the 16S rRNA genes. The similarity of communities determined by T-RFLP was examined as a function of incubation time at 3°C (A), temperature for 14 days (B), pressure (C), and sampling season (D) at 3°C for 14 days. The control (Cntrl) was filtered (pore size, 0.8 μm) untreated seawater at day 0.

The community similarity dendrogram for the effect of temperature revealed two main sample groupings; one group consisted of samples incubated at atmospheric pressure and the other group consisted of samples incubated at high pressure (Fig. 3B). The samples incubated at atmospheric pressure formed two distinct subclusters on the basis of the incubation temperature, while at 60 MPa the samples were less influenced by temperature, showing a higher community similarity (Fig. 3B).

Community analysis for the effect of pressure revealed three distinct sample clusters; one cluster included the control samples, a second cluster consisted of the samples incubated at 0.1 and 10 MPa, and a third cluster made up the samples incubated at 30, 60, and 80 MPa (Fig. 3C). The sample clustering correlated well with the species diversity values, with the higher similarity of the control sample and the samples incubated at high pressure reflecting higher species diversity (see Fig. S2A in the supplemental material). The higher pressures of 30 MPa, 60 MPa, and 80 MPa resulted in an increase in the community diversity, whereas low pressure (0.1 and 10 MPa) resulted in a decrease in diversity. The replicates showed high similarity to each other, and the sample collection date seemed to contribute to the community heterogeneity only for the samples incubated at high pressure (30, 60, and 80 MPa) (Fig. 3C).

Two distinct groupings were discernible in the dendrogram for seasonal effects on the variation in the community structure based on the sample collection date. One group included the samples incubated at atmospheric pressure, and the other group included the samples incubated at 60 MPa (Fig. 3D). The community structure in samples incubated at atmospheric pressure showed high heterogeneity, and the community structures in the two subclades (May-January and October-June) were highly dissimilar. The communities in samples incubated at high pressure were more similar; the communities in samples collected during May 2010 and October 2009 showed the highest similarity (Fig. 3D). Species diversity was lower for samples collected during May 2010 and October 2009 than for samples collected during June 2009 and January 2010, reflecting the clustering patterns (see Fig. S2D in the supplemental material).

Nonmetric multidimensional scaling (nMDS) analysis of the T-RFLP profiles based on the Bray-Curtis similarity indices (presence/absence and relative abundance) indicated significant differentiation of the community profiles in response to pressure (Fig. 4A). The profiles of the communities in samples incubated at 0.1 and 10 MPa appeared as separate groups in the bidimensional plot, while the profiles of the communities in samples incubated at 30, 60, and 80 MPa grouped together. The date of sample collection also had a significant effect on the variation in community structure (Fig. 4A). Analysis of variance of the sample parameters revealed that pressure accounted for most of the community shifts (P = 0.005) (Fig. 4B). Temperature and sample collection time accounted equally for the variance in T-RFLP community profiles observed (P = 0.02), while incubation time influenced the community changes to a smaller degree (P = 0.06) (Fig. 4B). Finally, a statistically significant interaction between pressure and incubation time was observed (Fig. 4B).

FIG 4.

Multivariance analysis of community T-RFLP profiles for all treated samples. (A) Nonmetric multidimensional scaling of T-RFLP profiles based on Bray-Curtis dissimilarities. Ellipses, standard deviations of the community structures; arrow, a correlation vector with significance (P < 0.1). (B) Results of multivariance analysis of variance for all the factors and their interactions between the communities whose 16S rRNA was analyzed by T-RFLP. Shown are mean square (MS), F, and P values for each factor.

Taxonomic diversity.

Next, the composition of the microbial communities present at 0.1 and 60 MPa was investigated in order to establish the effect of high pressure on community composition. Total DNA was extracted from seawater that had been incubated at 3°C for 14 days at 0.1 MPa or 60 MPa and used for 16S rRNA gene cloning and sequencing. Subsequent phylogenetic analyses revealed that at atmospheric pressure approximately 65% of the bacterial sequences recovered belonged to the Alphaproteobacteria, consisting of Rhodobacterales and one unclassified group (Fig. 5). The rest of the sequences were derived from the Epsilonproteobacteria (34%), consisting of Campylobacterales and one unclassified group, and the Betaproteobacteria (1%) of the order of Burkholderiales.

FIG 5.

(A) 16S rRNA gene sequence clone library analysis of coastal marine bacteria incubated at 0.1 MPa or 60 MPa for 14 days at 3°C in the dark. (B) Alphaproteobacteria-specific clone library analysis. *, P < 0.001.

In contrast, at high hydrostatic pressure, 87% of the bacterial sequences were classified as Alphaproteobacteria, consisting of three named orders (Rickettsiales SAR11 clade, Rhodospirillales, Rhodobacterales) and one unclassified group (Fig. 5). There were no sequences for Betaproteobacteria, while the relative abundance of epsilonproteobacterial sequences decreased significantly to only 1%. Moreover, at high pressure, 4% of the sequences were classified as Actinobacteria, 2% as Gammaproteobacteria, 1% as Flavobacteria, and the remaining 4% as unclassified bacteria. The results presented above suggest an overall increase in the diversity of the bacterial community at high pressure compared to that of the population at atmospheric pressure.

Calculations of the Shannon-Weaver diversity index (H′) for the two clone libraries also indicated higher community diversity at 60 MPa (H′ = 1.16) than at 0.1 MPa (H′ = 0.77). Thus, the results of the DNA-based T-RFLP and the 16S rRNA gene clone library analyses were in general agreement with one another. In contrast, when T-RFLP analyses were conducted with rRNA gene transcript preparations converted to cDNAs, the results were quite different. For the cDNA T-RFLP community profile at atmospheric pressure, the diversity index increased to 1.27, while at high pressure the diversity plummeted to 0.32 (see Fig. S2 in the supplemental material). At atmospheric pressure, the same major TRFs were present in both the cDNA and rRNA gene community profiles, while at high pressure there were only two TRFs present in the cDNA community profile (see Table S3 in the supplementary material).

Characterization of viable bacterial community members.

In order to identify some of the viable members of the bacterial community present in the seawater microcosm incubated for 2 weeks at 60 MPa and 3°C, following the experimental treatment, aliquots were plated on marine agar plates and incubated at 16°C (the in situ temperature) in the dark. The viable counts for samples incubated at atmospheric pressure were 2 orders of magnitude lower than the total cell counts, while the viable counts for the samples incubated at 60 MPa were 4 orders of magnitude lower (Fig. 6). There was no significant difference between the viable counts observed for samples incubated at 0.1 MPa or 10 MPa, while no viable cells were recovered from samples incubated at 80 MPa.

FIG 6.

Viable cell numbers (CFU ml⁻¹) of the microbial community recovered following incubation of seawater samples at 0.1 MPa, 10 MPa, 30 MPa, 60 MPa, and 80 MPa for 14 days at 3°C. Bar histograms indicate the mean value of four replicates, and the error bars indicate the standard deviations of the means. *, P < 0.05. ND, not detected.

The colonies recovered from the microcosms incubated at 60 MPa were found by 16S rRNA gene sequencing to belong to the Alphaproteobacteria, Gammaproteobacteria, and Actinobacteria. (More complete descriptions are provided in Table 1 and Fig. 7.) Intriguingly, several of the isolates were related to bacteria present in deep-sea settings. These environments included the Mariana Trench and the East Sea, deep-sea sediments in the Indian Ocean, and the gut of a deep-sea holothurian (strains AM3, AM4, and AM6, respectively). Another isolate, AM1, is a Sphingomonas sp. (Alphaproteobacteria) whose sequence is closely related to alphaproteobacterial sequences from Antarctic lakes. The similarity of some of the isolates to deep-sea bacteria, such as that of AM3 to the Mariana Trench isolate Micrococcus sp. strain JAM-AC11 (93% identity), could reflect an ability to withstand high pressure rather than grow at high pressure. Indeed, among all the isolates, only AM6 grew at pressures above 0.1 MPa (Fig. 8A). This piezotolerant strain grew similarly well at 0.1 and 20 MPa and progressively less well at 40 and 60 MPa. At atmospheric pressure, AM6 appeared rod shaped with an average cell diameter of 0.8 μm, while at increasing pressure, the cells reduced in size and assumed a coccoid morphology (Fig. 8B).

TABLE 1.

High-pressure-surviving isolates from samples incubated at 60 MPa for 14 days at 3°C in the dark^a

Isolate	Closest match	Sequence identity (%)	In silico TRF length (bp)	TRF length (bp)	Class	Order
AM1	Sphingomonas sp. strain Ant20	91.3	311	291	Alphaproteobacteria	Sphingomonadales
AM2	Brevundimonas sp. strain 626	96.3	55	38	Alphaproteobacteria	Caulobacterales
AM3	Micrococcus sp. strain GNUM-4	97.2	229	230	Actinobacteria	Actinomycetales
AM4	Kocuria marina KMM 3905	94.2	234	237	Actinobacteria	Actinomycetales
AM6	Halomonas sp. strain NK-W2-1	97.5	325	320	Gammaproteobacteria	Oceanospirillales

^aSequence identity was calculated using the EMBOSS Needle pairwise sequence alignment tool. Closest matches were identified using the Greengenes BLAST tool. The virtual digest positions were estimated on the basis of the predicted binding of primer 27F in relation to the E. coli 16S rRNA sequence.

FIG 7.

Phylogenetic relationships among the high-pressure-surviving isolates, closely related isolates, and selected piezophiles. The numbers at the nodes represent bootstrap values. The scale bar shows the average number of substitutions per site. *, piezophiles. GenBank accession numbers are given in parentheses.

FIG 8.

(A) Growth phenotypes of AM6, P. profundum SS9, and E. coli as a function of pressure. AM6 and P. profundum SS9 were incubated in MB at 16°C in the dark. E. coli was incubated in LB broth at 37°C in the dark. Each symbol indicates the mean value of at least three replicates, and the error bars indicate the standard deviations of the means. OD, optical density. (B) DAPI-stained microphotographs of isolate AM6, which survived at high pressure. AM6 was grown in MB at 0.1 MPa, 20 MPa, 40 MPa, and 60 MPa and 16°C in the dark.

Putative identification of bacterial OTUs.

In silico HaeIII digestion of the unique phylotypes from the 16S rRNA clone library and high-pressure-surviving isolates was performed to identify the major taxonomic units in the T-RFLP community (Table 1; see also Table S2 in the supplemental material). This approach was possible since both the 16S rRNA clone sequences and the T-RFLP community profiles were generated with the same 16S rRNA primers. There was good overlap between the in silico TRFs and the unique TRFs (15 operational taxonomic units [OTUs]) from the community analysis, with the difference between the predicted and observed sizes being 1 to 5 bp due to variability in the electrophoretic mobility of DNA fragments (44).

Closer examination of the changes in the relative abundance of certain TRFs revealed how pressure shaped the bacterial community composition (Fig. 9). The majority of the TRFs were identified as Alphaproteobacteria (113 bp, 193 bp, 221 bp, 219 bp, and 292 bp). The presence of pressure-tolerant species was indicated by pressure-dependent changes in abundance. The relative abundance of the majority of the putative alphaproteobacterial fragments increased significantly for samples incubated at pressures equal to or greater than 30 MPa (Fig. 9A, B, D, and E). The relative abundance of the putative alphaproteobacterial fragment of 221 bp, identified as a member of the Rhodospirillaceae, decreased to undetected levels for samples incubated at pressures higher than 30 MPa (Fig. 9C). A similar pressure-sensitive response was also observed for fragments of 324 bp, identified as Gammaproteobacteria, and 410 bp, identified as Bacteroidetes (Fig. 9F and H). However, contrary to the putative Bacteroidetes fragment that was undetectable at high pressures above 30 MPa, at 80 MPa the gammaproteobacterial fragment could still be detected at levels comparable to those for the control samples (Fig. 9F).

FIG 9.

Univariance analysis and putative identification of selected terminal restriction fragments from samples incubated for 14 days at 3°C. Bar histograms indicate the mean values of four replicates, and the error bars indicate the standard deviations of the means. The control (Cntrl) was untreated filtered (pore size, 0.8 μm) seawater at day 0.

Interestingly, the sizes of the TRFs produced by two of the isolated piezotolerant strains, AM2 and AM6, matched the sizes of the TRFs detected in the rRNA community profiles at high pressure. AM2 produced a 38-bp fragment and AM6 produced a 320-bp fragment when their 16S rRNA genes were analyzed via T-RFLP (Table 1). A 38-bp TRF accounted for 92% of the active community abundance at 60 MPa and a 320-bp TRF accounted for 8% (see Table S3 in the supplementary material).

DISCUSSION

In this study, the impact of high pressure on a shallow-water coastal microbial community was assessed as a function of the magnitude and duration of the pressure applied. The conditions examined differed in several fundamental ways from those employed in related studies (21–23). This generally included (i) the application of a greater range of pressures, (ii) the omission of nutrients other than those present in the collected seawater, and (iii) rapid pressurization rather than gradual pressurization at a rate microbes associated with sinking organic particles would be expected to encounter.

Abundance and morphology.

High-pressure treatment of samples from shallow water resulted in a dramatic decrease in total cell numbers following 8 days of treatment. Variable growth rates (0.2 to 3 days) combined with an oligotrophic environment, due to preincubation and filtration (pore size, 0.8 μm), could account for the observed 8-day lag phase (45). Even though temperature had little effect on cell abundance at 60 MPa, samples collected in January 2010 responded to pressure changes with a drop in total cell numbers that was less abrupt. Seasonal variability in cell abundance could be attributed to lower coastal water temperatures in January 2010 (14 to 15°C) compared to those in October 2009 (16 to 21°C). Bacteria share common adaptation responses to low temperature and high pressure, such as changes in membrane composition, including an increase in the fraction of unsaturated fatty acids (46–48). Shifting the temperature of a soil microbial community from 25 to 10°C resulted in higher levels of unsaturated fatty acid production over the course of 14 days (49). Higher cell abundance for samples collected in January 2010 could reflect seasonal changes to the membrane composition that confer pressure tolerance.

Previous studies have established that shallow-water microbes respond to elevated pressure by filamentation (19, 40). Under nutrient-replete, high-pressure conditions that still enable biomass accumulation (up to ∼40 MPa), E. coli cells become filamentous due to dissociation of the cell division cytoskeletal framework and/or the induction of the SOS DNA damage response (5, 50). High-pressure treatment of shallow-water microbes in a high-nutrient medium results in elongated cells (51, 52). However, elongated cells are not observed at pressures precluding biomass accumulation. Increasing the pressure to 60 MPa reverses the filamentous morphology of E. coli by encouraging cell separation (53), an observation that is also generally consistent with the observations for the coastal samples examined here with or without added nutrients. The absence of a filamentous stage for AM6 at increasing pressure could signify the ability of AM6 to withstand changes in pressure and remain viable after a prolonged incubation at 60 MPa. Another possibility is that elevated pressure is inducing the onset of stationary phase. Some bacteria, including E. coli, switch from rods to cocci as they transition to stationary phase due to reductive division (54).

Community analysis.

The results of phylogenetic analysis of the predominant taxa at atmospheric pressure agree with those of previous community analyses of coastal water collected off the pier of the Scripps Institution of Oceanography (55, 56). Our results suggest an increase in the diversity of the Alphaproteobacteria at 60 MPa. Some of these differences are consistent with depth-related differences in the bacterial composition of deep (nonreducing zone) versus shallow water masses (11). The increase in the relative abundance of flavobacteria observed at 60 MPa was also reported in diatom detritus sinking experiments at 15 MPa (22). Pressure-resistant spores of surface-derived Clostridium spp. have been recovered from sediments at depths of 7,000 m, while mesophilic actinomycetes have been recovered from sediments from an ∼10,900-m depth within the Challenger Deep (57, 58).

Surface marine microbes can be divided in two broad groups: the abundant microbes with little metabolic plasticity and the microbes with a low abundance that can persist under unfavorable conditions due to their lack of specialization (59). Changes in the relative abundance of the rare microbes usually reflect changes in the environment, such as nutrient availability or, in the present study, even pressure. Our findings suggest that when the environmental conditions changed, the rare microbes became more abundant, as reflected in the increase of the overall diversity in samples incubated at high pressure. They were able to outcompete other community members due to their ability to tolerate increases in pressure.

Interestingly, there was a dramatic shift in the diversity for the active community, as reflected by the rRNA T-RFLP profiles at 60 MPa. The robustness of DNA and the presence of nonmetabolically active cells could contribute to an overestimation of the pressure-resistant phylotypes, while the active bacteria represented only a small fraction of the total community (60, 61). The sizes of the TRFs recorded for the active community at 60 MPa matched the sizes of the TRFs produced for two out of the five characterized high-pressure-surviving isolates, confirming that they were members of the active community at high pressure and validating the ability of AM2 and AM6 to tolerate high pressure.

Why should either piezoresistant or piezotolerant microbes that are related to deep-sea microbes exist in a coastal environment? One possibility is that they originate from the deep sea and have been introduced to shallow waters by a mixing process, such as upwelling, a process well documented to occur in Southern California (62). Ocean currents contribute to shaping the composition of marine microbial communities (63). The idea that high-pressure-adapted microbes could exist in a shallow-water setting is a variation on the theme expressed by Baas Becking: “everything is everywhere, but, the environment selects” (64). Resistance to high pressure is easily selected for in many microbes at pressures up to the gigapascal range (65). Given the close phylogenetic relationship between low-temperature piezophilic microbes and some low-temperature and presumably piezosensitive polar microbes, it might even be possible for psychrotolerant coastal microbes to acquire mutations conferring improved growth fitness at high pressure (66).

Pressure as an important parameter.

Our results highlight the importance of pressure as an environmental parameter. Under the conditions employed, pressure was the most important parameter in shaping the microbial community structure. Increasing the pressure 100-fold caused a transient community response, while a 300-fold increase caused substantial changes to the microbial community. The close match between the community profiles at 30, 60, and 80 MPa determined by T-RFLP suggests that increasing the pressure to pressures above 30 MPa did not substantially alter the community structure any further. The significant effect of high pressure at 30 MPa was also reflected in the low levels of cell viability, while at 80 MPa, the effects of pressure on the microbial community were fatal. Pressure exerts a strong selection force and should be accounted for in studies addressing environments with pressures of 10 MPa or higher, since moderate changes to pressure (0.1 to 10 MPa and 30 MPa) cause significant variation to the microbial community structure.

Even though hydrostatic pressure is an important physical parameter that shapes the biogeography of life, there are few studies that have addressed the response of shallow-water marine microbes to high pressure (18–23). The issue is significant, given that a variety of physical and biological processes can introduce large numbers of surface-water and shallow-sediment microbes to great depth, as has been convincingly demonstrated for selected sporeformers and phytoplankton (14, 57, 58). Thus, shallow-water microbial community activity and survival as a function of pressure are important to understanding the nature of all microbial life (allochthonous and autochthonous) at depth, in addition to providing an important foundation from which to view the distribution and properties of piezophiles.