Front. Microbiol.Frontiers in MicrobiologyFront. Microbiol.1664-302XFrontiers Media S.A.10.3389/fmicb.2022.867340MicrobiologyOriginal ResearchCharacterizing the Piezosphere: The Effects of Decompression on Microbial Growth DynamicsCarioAnaïs1*†‡OliverGina C.1†‡RogersKaryn L.12*1Department of Earth and Environmental Sciences, Rensselaer Polytechnic Institute, Troy, NY, United States2Rensselaer Astrobiology Research and Education Center, Rensselaer Polytechnic Institute, Troy, NY, United States
Edited by: Andreas Teske, University of North Carolina at Chapel Hill, United States
Reviewed by: Philippe M. Oger, UMR5240 Microbiologie, Adaptation et Pathogenie (MAP), France; Aude Picard, University of Nevada, Las Vegas, United States
*Correspondence: Anaïs Cario, anais.cario@cnrs.frKaryn L. Rogers, rogerk5@rpi.edu
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The extent to which the full diversity of the subsurface microbiome can be captured via cultivation is likely hindered by the inevitable loss of cellular viability from decompression during sampling, enrichment, and isolation. Furthermore, the pressure tolerance of previously isolated strains that span surface and subsurface ecosystems can shed light into microbial activity and pressure adaptation in these transition zones. However, assessments of the effects of elevated pressure on the physiology of piezotolerant and piezosensitive species may be biased by high-pressure enrichment techniques. Here, we compared two high-pressure cultivation techniques—one that requires decompression of the whole cultures during sampling and one that employs the previously described isobaric PUSH devices—to explore the effects of repeated decompression during incubations performed to characterize isolates from deep environments. Two model sulfate-reducing prokaryotes were used to test the effects of decompression/repressurization cycles on growth rates, cell yields, and pressure tolerance. The mesophilic bacterium Desulfovibrio salexigens was cultivated from 0.1 to 50 MPa, and the hyperthermophilic archaeon Archaeoglobus fulgidus was tested from 0.1 to 98 MPa. For both cultivation methods, D. salexigens showed exponential growth up to 20 MPa, but faster growth rates were observed for isobaric cultivation. Furthermore, at 30 MPa minor growth was observed in D. salexigens cultures only for isobaric conditions. Isobaric conditions also extended exponential growth of A. fulgidus to 60 MPa, compared to 50 MPa when cultures were decompressed during subsampling. For both strains, growth rates and cell yields decreased with increasing pressures, and the most pronounced effects of decompression were observed at the higher end of the pressure ranges. These results highlight that repeated decompression can have a significant negative impact on cell viability, suggesting that decompression tolerance may depend on habitat depth. Furthermore, sampling, enrichment, and cultivation in isobaric devices is critical not only to explore the portion of the deep biosphere that is sensitive to decompression, but also to better characterize the pressure limits and growth characteristics of piezotolerant and piezosensitive species that span surface and subsurface ecosystems.
Most of the bacterial and archaeal biomass on Earth is in deep-sea and subsurface environments at elevated pressures (Whitman et al., 1998; Kallmeyer et al., 2012; Parkes et al., 2014; Bar-On et al., 2018). These microorganisms have been shown to be well-adapted to the elevated pressures of their natural habitats (e.g., Somero, 1992; Allen and Bartlett, 2002; Simonato et al., 2006; Jebbar et al., 2015; Peoples and Bartlett 2017). Despite this, pressure seems to be one of the least explored parameters for microbial growth, as relatively few microorganisms from these deep environments have been isolated and/or grown under in situ pressures. To date, fewer than 100 species have been reported to be piezotolerant or piezophilic (Picard and Daniel, 2013; Jebbar et al., 2015; Cario et al., 2019; Oliver et al., 2020; Alain et al., 2021; Courtine et al., 2021; Li et al., 2021; Yu et al., 2021; Zhang et al., 2022), and few obligate piezophiles have been identified (Bartlett, 2002; Zeng et al., 2009). Therefore, our knowledge of active microbial species, their physiological and metabolic potential, and community diversity in these deep environments is limited.
Exploration of high-pressure life is limited by the difficulty in both sampling from high-pressure environments and replicating those pressure conditions in the laboratory during cultivation, isolation, and characterization. For obligate piezophiles, high-pressure sampling and cultivation is the only route to isolate novel species (Yayanos and Dietz, 1983; Jannasch and Wirsen, 1984). However, facultative piezophiles and piezotolerant microorganisms can often withstand lower sampling and transfer pressures (Jannasch and Wirsen, 1984). In these cases, the total change in pressure, duration of decompression, and number of subsequent decompression/repressurization cycles can impact the successful isolation of piezophiles (Yayanos, 2001; Park and Clark, 2002; Peoples and Bartlett, 2017). The pressure condition of cultivation experiments may also impose a selection bias on enrichment and isolation experiments that favor species more tolerant to decompression or lower growth pressures (Jannasch et al., 1992; Yayanos, 1995; Grossart and Gust, 2009). Furthermore, Park and Clark (2002) showed that rates of decompression could also impact microbial survival during decompression. For example, accelerated rates of decompression (26 MPa/s) caused the piezophile, Methanococcus jannaschii cells to rupture while slower rates of decompression (5.2 MPa/min) over the same pressure range increased viable cell yields (Park and Clark, 2002). Additionally, many piezophiles recovered from the intestinal systems of deep-sea macro fauna (Yayanos et al., 1979; Deming et al., 1981; Jannasch and Wirsen, 1984), show a greater tolerance to sample decompression, but the loss of species from decompression has yet to be quantified and any correlation with habitat depth has yet to be explored (Yayanos, 1978).
Deep biosphere species are often exposed to decompression during sample recovery, as well as during transfer, enrichment, and isolation. Therefore, significant effort has gone into developing pressure-retaining vessels to sample from the deep ocean habitats and carry out enrichment and isolation experiments without decompression (Tabor and Colwell, 1976; Jannasch and Wirsen, 1977; Yayanos, 1977; Cahet et al., 1990; Bianchi et al., 1999; Tamburini et al., 2003; Kato et al., 2008; McNichol et al., 2016; Cario et al., 2019). Such variable volume, floating piston devices can maintain elevated pressure during subsampling, inoculation, and/or transfer (Bianchi et al., 1999; Tamburini et al., 2003; Garel et al., 2019). Recent application of these new technologies has confirmed that higher rates of microbial activity and cell growth are achieved in incubation studies of bathypelagic samples maintained at in situ pressures without decompression, compared to parallel, decompressed, ambient pressure experiments (Tamburini, 2006; Tamburini et al., 2013; Garel et al., 2019, 2021). These high-pressure incubation experiments emphasize the need to study deep-sea microbes under in situ pressure conditions in order to accurately quantify deep-sea prokaryotic activity.
For enrichments and isolation, common methods involve growth in syringes, plastic bulbs, or heat-sealed plastic bags, in static pressure vessels (reviewed in Yayanos, 2001). However, subsampling to monitor cell growth over time requires decompression of the entire system followed by repressurization for continued cultivation. Inoculating such devices for high-pressure enrichment experiments also usually requires ambient pressure conditions. Other alternatives to study microorganisms under in situ pressure conditions include (i) the implementation of optic windows in the high-pressure vessels to monitor growth and motility (Maldonado et al., 2016; Garel et al., 2019); (ii) using high-pressure capillaries or other high-pressure cells coupled with microscopy to study molecular behaviors under extreme conditions with specific high-pressure cell using fluorescence microscopy (Raber et al., 2006; Usui et al., 2012; Patra et al., 2017; Bourges et al., 2020); and (iii) application of spectroscopy techniques to characterize microbial metabolism under high-pressure conditions (Kato and Fujisawa, 1998; Picard et al., 2007, 2015; Peters et al., 2014; Martinez et al., 2016; Osman et al., 2021). These alternatives are promising but require both specific and costly equipment and expertise to perform high-pressure in situ monitoring, and are not-yet widespread.
Microbial Strains and Growth MediumSelection of Target Strains for HT-HHP Growth
The choice of D. salexigens and A. fulgidus allowed for a comparison between two strains with similar metabolic strategies (heterotrophic sulfate reduction) over a range of temperatures (D. salexigens is a mesophile, while A. fulgidus is a thermophile) for two genera that are ubiquitous in subsurface environments and also represent both prokaryote. Each of these strains have had previous indications of growth at elevated pressure, belong to genera with other piezotolerant/piezophilic species, and also have been identified in high-pressure ecosystems. For example, a previous study, based on sulfide production measurements, showed that D. salexigens was active up to 5 MPa (Bale et al., 1997), and three other Desulfovibrio species have been isolated from subsurface environments (D. profundus, D. hydrothermalis, and D. piezophiles; Bale et al., 1997; Alazard et al., 2003; Khelaifia et al., 2011). While A. fulgidus type strain VC-16 was isolated from a shallow marine vent (Stetter et al., 1987, 1988), this strain has been identified in a number of deep-sea and deep subsurface environments (1–4 km and ~10–40 MPa; Stetter et al., 1987; Beeder et al., 1994; L'Haridon et al., 1995; Nakagawa et al., 2005; Fardeau et al., 2009). Additional species within the genus Archaeoglobus have been isolated from deep-sea environments (e.g., Archaeoglobus veneficus was isolated from the Mid Atlantic Ridge at 3.5 km depth, Huber et al., 1997), and archaeal sequences belonging to Archaeoglobaceae have been identified at the Mid-Cayman Rise, the deepest known hydrothermal system that reaches 4.96 km depths (~50 MPa; Reveillaud et al., 2016). Further, we have previously reported high-pressure growth of A. fulgidus (Oliver et al., 2020), showing that this strain is tolerant up to ~30–40 MPa (with decompression), which is consistent with the depths and pressures of deep-sea and deep subsurface environments where it has been identified.
Growth Conditions for <italic>Desulfovibrio salexigens</italic> and <italic>Archaeoglobus fulgidus</italic>
Desulfovibrio salexigens (DSM 2638), a marine sulfate-reducing bacterium (Postgate and Campbell, 1966) was obtained from the Deutsche Sammlung von Mikroorganismen und Zellkulturen (Germany) and grown anaerobically in DSMZ medium 163 (Solution A: NaCl 25 g L−1, K2HPO4 0.5 g L−1, NH4Cl 1.0 g L−1, Na2SO4 1.0 g L−1, CaCl2.2H2O 2.0 g L−1, MgSO4.7H2O 2.0 g L−1, Na-DL-lactate 2.0 g L−1, Yeast Extract 1.0 g L−1, resazurin solution (0.1% w/v) 0.5 ml L−1, distilled water 980 ml; Solution B: FeSO4.7H2O 0.5 g/10 ml and Solution C: Na-thioglycolate 0.1 g/10 ml, ascorbic acid 0.1 g/10 ml), similar to Postgate and Campbell (1966), with the exception that the media was enriched in calcium (1.5 times) and reduced in iron (1:100). Solutions B and C were added to solution A under N2 and the pH was adjusted to 7.8 with NaOH.
Archaeoglobus fulgidus strain VC-16 (DSM 4304) was obtained from the Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH (DSMZ, Braunschweig, Germany). Archaeoglobus fulgidus VC-16 is an anaerobic, hyperthermophilic sulfate-reducing archaeon. Here, A. fulgidus was grown chemolithoheterotrophically in a lactate–sulfate-rich medium. The composition of the culture medium followed Hartzell and Reed (2006) and sterile anaerobic conditions were maintained following Balch et al. (1979). The medium was reduced prior to inoculation by adding Na2S.9H2O to a final concentration of 0.1% prior to inoculation (Cario et al., 2016).
Desulfovibrio salexigens cultivations with or without decompression (static pressure vessels, and PUSH vessels, respectively) were tested at 0.1, 10, 20, 30, 40, and 50 MPa. Subsamples were taken at ~12 h intervals for 72 h to obtain a robust growth curve. For the higher pressures (30–50 MPa), a cell count after 150 h of growth was performed to enumerate cell density (death or survival) according to the pressure cultivation technique. Cell recovery experiments were performed for this strain in order to evaluate the impact of elevated pressure on cell growth recovery. The protocol and the obtained results are detailed in Table 1. Archaeoglobus fulgidus VC-16 was cultivated from 0.1 to 98 MPa in ~10 MPa increments at 83°C in PUSH vessels and in static pressure vessels. Six to eight subsamples were taken at regular time intervals for standard growth curves. Archaeoglobus fulgidus cells grown in the glass syringes were decompressed and repressurized a maximum of nine times throughout each batch culture experiment.
The percentage of D. salexigens cell recovery after 36 h of growth at different elevated pressures, with or without decompression steps, and transfer at ambient pressure.
Cultivating pressures
Percentage of growth recovery after transfer at ambient pressure
Dilutions
After 24 h of growth
After 48 h of growth
Decompressed
No decompressed
Decompressed
No decompressed
%
SD
%
SD
%
SD
%
SD
10 MPa
1
50.7
1.7
90
2.2
76.7
12.5
99
0.5
10
81.8
5.1
82
1.7
84.8
8.3
97
1.2
100
52.5
17.2
81
3.6
95.5
3.9
99
0.8
20 MPa
1
20.2
2.2
76
2.8
69.1
14.8
98
1.1
10
12.6
3.5
68
1.7
78.3
18.8
99
0.2
100
10.2
3.7
71
5.2
64.2
15.1
99
0.8
30 MPa
1
NG
50
4.3
NG
89
2.5
10
48
2.6
91
3.1
100
55
3.8
85
5.8
40 MPa
1
NG
37
6.2
NG
80
5.2
10
39
2.5
78
3.9
100
31
4.8
81
8.7
50 MPa
1
NG
25
5.7
NG
78
10.2
10
22
7.2
72
8.9
100
17
12.3
70
15.2
The cell counts were measured after 24 and 48 h of growth at ambient pressure after the transfer from high-pressure to ambient pressure. For each pressure condition (P), one non-pressurized culture (ambient pressure) was used as a control (U). The average ratio of the growth recovery (U/P) was calculated for each dilution series and was performed at least in triplicate. SD, standard deviation and NG, no growth was observed. Significant differences were determined by Student’s t-test (value of p < 0.01).
HT-HHP Culture ExperimentsPre-cultures and Inoculation
For triplicate HT-HHP culture experiments with and without sample decompression, the sterile media and pre-cultures were prepared for four PUSH vessels and four syringes incubated in static pressure vessels, corresponding to three inoculated culture replicates and one uninoculated experiment to serve as a negative control. Each strain was first grown from a frozen stock (−80°C) and transferred into sterile anaerobic growth medium. Exponential growth of this culture was used as inoculum for three independent pre-cultures. After reaching logarithmic-phase growth, each of these pre-cultures was used to inoculate fresh sterile growth medium in three serum bottles. Immediately following inoculation, 8–10 ml from each inoculated serum bottle were transferred into sterile syringes and the remaining 45–47 ml were transferred into each PUSH vessel. The D. salexigens culture started at 1.106 cells/ml and incubations were carried out in static pressure vessels and PUSH vessels at 30°C. Inoculation and transfer of A. fulgidus cultures were carried out in the anaerobic chamber in pre-heated PUSH vessels. Starting cell concentrations for A. fulgidus experiments were ~6.7 × 106 cells/ml, and incubations were carried out at 83°C.
PUSH Vessel Preparation, Pressurization, and Subsampling
For each batch culture HT-HHP experiment, four PUSH vessels were sterilized, assembled, and pre-heated before inoculation and transfer (Supplementary Information 1, Supplementary Figure S1). After pre-assembly, each PUSH was individually wrapped in an insulated temperature-controlled system with thermocouple and heating jacket to 30°C and 83°C, for D. salexigens and A. fulgidus, respectively (Supplementary Information 1, Supplementary Figure S1B). Once pre-heated, all four vessels were transferred with their respective temperature-control systems into an anaerobic chamber (Bactron Shellab), and each temperature-control system was plugged into the DC to AC power converter allowing for continual vessel heating throughout the anaerobic inoculation process. In the anaerobic chamber, the PEEK reservoirs of each of the four PUSH vessels were filled with ~45–47 ml of pre-inoculated growth medium (triplicate) or sterile growth medium (Supplementary Information 1, Supplementary Figure S1C). The PEEK reservoir screw cap and lid was then closed completely and the valves were closed while in the anaerobic chamber. The vessels were then transported out of the anaerobic chamber for pressurization.
Each PUSH vessel was individually pressurized to the target pressure with a hydraulic screw pump with an in-line pressure gauge connected to the PEEK piston valve (Supplementary Information 1, Supplementary Figure S1D). Once pressurized, an initial 0.5 ml subsample was taken and fixed in 2.5% glutaraldehyde from each PUSH vessel using the decompression line (see below). Initial sampling and pressurization of the all of the PUSH vessels and syringes were done within 1 h of inoculation in the anaerobic chamber. To assure pressure stability, potential pressure loss from vessel leakage was closely monitored for the first 3 h after inoculation. Up to nine subsamples were obtained from each PUSH vessel for every HT-HHP batch culture growth experiment. For subsampling, the average decompression rate was of 15–25 MPa/min (Supplementary Information 1, Supplementary Figure S1E). The first 3 ml of medium sampled were discarded as waste from flushing the decompression line before taking a 0.5 ml aliquot sample for enumeration. A maximum of 10% pressure loss occurred during subsampling, but in all cases the pressure was rapidly re-established. The decompression line was cleaned with 70% ethanol and ultrapure water (18.2 MΩ) before and after subsampling each PUSH vessel.
Syringe Preparation, Pressurization, and Subsampling
Cell enumeration was estimated by direct counts from fixed cells [2.5% (v/v) glutaraldehyde] in a Thoma-chamber (depth: 0.02 mm; Brand, Wertheim, Germany) using a light microscope (model XM: Olympus) under 80x magnification (e.g., Huber et al., 1989; Hei and Clark, 1994; Blöchl et al., 1997; Cario et al., 2016). Specific growth rates (μ-hr) were calculated from linear regressions of the exponential portion of the growth curves from triplicate experiments using the LINEST function in Excel. Error bars indicate the standard error from linear regressions of triplicate experiments. Cell densities quantified at ~36 h for A. fulgidus and 48 h for D. salexigens were used to evaluate the effects of different cultivation conditions on overall cell yields. As some pressure conditions led to cell death, maximum cell densities could not be used for comparison. Error bars indicate SD from triplicate experiments. Significant differences were determined by Student’s t-test (value of p < 0.01).
Results<italic>Desulfovibrio salexigens</italic> Growth at Elevated Pressure
Elevated pressure experiments were performed from ambient (0.1 MPa) up to 50 MPa for D. salexigens using both static pressure vessels (i.e., with cyclic decompression) and variable volume PUSH vessels (i.e., isobaric) for experiments that lasted 76–150 h. Growth curves, growth rates, and cell density data are summarized in Figure 1, Supplementary Table S1. At some elevated pressures, cell numbers decreased with time (e.g., cell death). To account for both cell growth and cell death at different conditions, cell densities are compared at 48 h over the entire pressure range of the experiments (Figure 1D).
(A)Desulfovibrio salexigens growth curves in plastic syringes with multiple sample decompression and (B)D. salexigens growth curves in the pressurized underwater sample handler (PUSH) vessels without multiple sample decompression from 0.1 to 50 MPa. (C)Desulfovibrio salexigens specific growth rates in the PUSH vessels (orange squares) and in syringes (blue circles). (D)Desulfovibrio salexigens maximum log cell densities in the PUSH vessels (orange squares) and in syringes (blue circles), open orange squares and blue circle indicate maximum cell densities in samples with no observed growth or observed cell densities lower than the average initial cell densities (dashed lines in D). Error bars are the SDs from the average of triplicate experiments. Significant differences were determined by Student’s t-test (value of p < 0.01).
Exponential growth of D. salexigens was observed up to 20 MPa both in the static and PUSH vessels (Figures 1A,B). The highest growth rates were observed at ambient pressure (0.1 MPa) and 30°C, and were nearly identical for cultivation in serum bottles and the PUSH vessels (0.19 ± 0.005 h−1 and 0.19 ± 0.004 h−1, respectively). The same growth rate was confirmed with D. salexigens incubation, performed in syringes, at ambient pressure and 30°C (data not shown). Overall growth rates decreased with pressure, but declining growth rates were less pronounced for isobaric enrichments (Figure 1C; Supplementary Table S1). For example, at 10 MPa the growth rate for isobaric cultivation was 0.17 ± 0.2 h−1, ~10.5% lower than optimal conditions, while cultivation with decompression had a ~36.8% drop in growth rate to 0.12 ± 0.1 h−1. Furthermore, the upper pressure limit for growth was higher in isobaric experiments, with slow growth observed at 30 MPa in PUSH vessels (0.02 h−1), compared to distinct cell death in decompressed experiments (Figures 1A–C).
While differences in growth rates were notable between the two cultivation techniques at moderate pressures (0.1–20 MPa), there were negligible effects on cell yields in this pressure range (Figure 1D; Supplementary Table S1). However, at supra-optimal pressures (≥30 MPa for D. salexigens), differences in cell densities (quantified at 48 h) were more significant. For cultivation experiments that included cyclic decompression, cell numbers decreased markedly with time at 30, 40, and 50 MPa, indicating cell death. Conversely, isobaric cultivation at these pressures led to only minor changes in cell density with time. For example, at 40 MPa, cell density decreased from an initial value of 106.30–105.78 after 48 h, while the isobaric culture had a cell density of 106.34 after 48 h and no decompression (Figure 1D). When considering changes in cell yields, decompression had a more significant impact at supra-optimal pressures, inducing cell death, as opposed to isobaric conditions, which only limited growth.
To further explore the impact of decompression on D. salexigens cell viability, cells were incubated at various elevated pressures (10–50 MPa) for 36 h, with or without decompression. They were then transferred to ambient pressure, incubated in fresh growth medium at several dilutions (1:1, 1:10, and 1:100, v/v), and cell recovery was estimated after 24 and 48 h of growth. Because the syringe cultivation technique requires decompression of the high-pressure vessels for subsampling, each vessel was subjected to six decompression/repressurization cycles with a minimum of 4 h between cycles. For both static pressure vessels and PUSH vessels, cells from the 10 and 20 MPa growth conditions were able to recover the optimal pressure cell density (see Table 1). However, when the cells were incubated at higher pressure (≥30 MPa) with cyclic decompression the cells were not able to recover and did not grow after 24 or 48 h of transfer at ambient pressure (nor after 1 week of incubation, data not shown). In comparison, cells exposed to 30–50 MPa in the PUSH without decompression remained viable, though were slower to recover as pressures increased. After transfer to and growth at ambient pressure (0.1 MPa) for 48 h, cell densities were only slightly lower (90% for 30 MPa and 70% for 50 MPa) than those observed for cells never exposed to elevated pressure (Table 1).
<italic>Archaeoglobus fulgidus</italic> Growth at Elevated Pressure
Archaeoglobus fulgidus VC-16 (type strain) was grown in batch cultivation experiments at 83°C over a range of pressures (0.1–98 MPa, at ~10 MPa increments) in PUSH vessels (isobaric) and in static pressure vessels (cyclic decompression) and cell densities were monitored up to 115 h. Growth curves, growth rates, and cell density data are summarized in Figure 2; Supplementary Table S1. To assess the impact of decompression on cell yields even in the cases of cell death, cell densities at 36 h are reported over the entire pressure range (Figure 2D). This time aligns with the generally faster growth of this strain, captures the exponential phase at lower pressures, and accurately represents cell death at elevated pressures.
(A)Archaeoglobus fulgidus growth curves in glass syringes with multiple sample decompression and (B)A. fulgidus growth curves in the PUSH vessels without multiple sample decompression from 0.1 to 98 MPa. (C)Archaeoglobus fulgidus specific growth rates in the PUSH vessels (orange squares) and in syringes (blue circles). (D)Archaeoglobus fulgidus maximum log cell densities in the PUSH vessels (orange squares) and in syringes (blue circles), open orange squares and blue circle indicate maximum cell densities in samples with no observed growth or observed cell densities lower than the average initial cell densities (dashed lines in D). Error bars are the SDs from the average of triplicate experiments. Significant differences were determined by Student’s t-test (value of p < 0.01).
Based on the growth curves shown in Figures 2A,B, A. fulgidus growth rates were calculated for pressures between 0.1 and 60 MPa (Figure 2C). Overall, A. fulgidus growth characteristics showed trends with pressure similar to those observed for D. salexigens. Exponential growth was observed in the PUSH vessels at 60 MPa until ~29 h, after which cell numbers decreased. In static pressure vessels at this pressure, only minor increases in cell density (1.34 ± 0.11 × 107 cells/ml) relative to initial values (6.03 × 106 ± 1.02 cells/ml) were observed, but were not consistent with a true exponential growth phase (Figure 2A). Growth was generally faster with higher maximum cell densities (with the exception at 10 MPa) in PUSH vessels compared to static pressure vessels, especially at the higher end of the pressure range (Figures 2C,D). The fastest growth rate in the PUSH vessels was measured at 0.1 MPa (0.326 ± 0.017 μ-hr), though growth rates were only slightly slower from 10 to 30 MPa (0.258 ± 0.026–0.302 ± 0.011 μ-hr). The largest difference in growth rates between the two cultivation methods was observed at 30 and 40 MPa (15.3% and 44%, respectively). Overall, growth rates indicate that A. fulgidus VC-16 is piezosensitive with a maximum exponential growth pressure of 60 MPa in isobaric conditions and 50 MPa for cultivation that included cyclic decompression.
Similar to the mesophilic strain, D. salexigens, the two cultivation techniques had a more significant impact on A. fulgidus cell density in the supra-optimal pressure range (≥50 MPa), while density was largely unaffected at the lower pressures. For example, from 0.1 to 40 MPa the largest difference in density at ~36 h was at 30 MPa (108.66, isobaric vs. 108.56, decompressed). Comparing isobaric growth to cultivation with cyclic decompression shows that the largest disparities in cell density (measured at ~36 h) were observed at 60 MPa when growth was observed and at 90 MPa when only cell death was recorded. Again, these data suggest that cycles of decompressions can accelerate loss of cell viability. Loss of cell viability was also tested via high-pressure incubation experiments in which A. fulgidus cells were incubated at 80 MPa for 115 h and subsequently transferred (10% v/v) to fresh growth medium and incubated at 0.1 MPa and 83°C. Ambient pressure cultures were monitored visually and no growth was observed after a week of incubation.
Pressure-Induced Effects on Cell Mobility and Morphology
We also observed significant effects of elevated pressure on the mobility and the morphology of D. salexigens cells using a light microscope (Supplementary Figure S3). Under optimal pressure conditions (i.e., ambient pressure, 0.1 MPa), the cells were highly motile and formed 3–4 μm vibrios and up to 20 μm cell long when reaching the late exponential and stationary phases (Supplementary Figure S3). Under high-pressure cultivation conditions (10 and 20 MPa) and subsequent sampling, the cells were barely motile and formed short vibrios (2–3 μm). After longer incubation times they also formed filamentous cells (Supplementary Figure S1). At higher cultivation pressures without decompression steps (30–50 MPa) and subsequent sampling, the cells were also immotile. When they were transferred for growth at ambient pressure, the cell division appeared altered with formation of cells arranged in chains (Supplementary Figure S4). Irregular and elongated cell morphologies were previously observed in A. fulgidus at elevated pressures (Oliver et al., 2020). This morphological feature has been reported in several studies, where elevated pressure inhibited the cell division protein FtsZ (e.g., Ishii et al., 2004; Abe et al., 2013). Morphological changes are a common stress response (e.g., Zobell and Oppenheimer, 1950; Donaldson et al., 1989) and might be a global stress response to high-pressure (e.g., Bidle and Bartlett, 1999; Aertsen et al., 2004).
Additionally, for elevated pressure conditions (10–50 MPa) we noticed the presence of cyst-like cells (Supplementary Figure S3), a constitutive dormancy behavior characteristic of some non-spore forming bacteria (Suzina et al., 2004). As pressure increased from 10 to 50 MPa, we observed an increasing abundance (approximately 2-fold) of such pleiomorphic cells (Supplementary Figure S5). However, after transferring these cultures to ambient pressure, we observed the release of cells resembling the development of vegetative cells, suggesting that these structures could be cysts (Supplementary Figure S4). The formation of various types of structures, such as endospores, exospores, or cysts, is previously recognized survival strategy of many bacteria under unfavorable conditions induce (Sudo and Dworkin, 1973). The pressures at which we observed changes morphologies and structures are consistent with the growth data that indicated the onset global stress, as indicated by slower growth rates and decreasing cell numbers. Interestingly, among the dissimilatory sulfate-reducing bacteria, the genus Desulfovibrio was characterized as non-sporulating compared to the closely related genus Desulfotomaculum (Postgate and Campbell, 1966). Such behavior might help this species to cope with extreme growth conditions, such as elevated pressure conditions, and might be an important contribution for microbial dissemination in various environments. These specific structures deserve more investigations to determine the developmental steps and the ultrastructural properties of this species when growing under unfavorable growth conditions.
Discussion
A majority of the Earth’s biosphere thrives in high-pressure, subsurface environments (Fang et al., 2010; Oger and Jebbar, 2010; Picard and Daniel, 2013), and like many natural ecosystems, studies of subsurface diversity reveal a vast number of uncultured species, often only identified through the presence of their genetic material. The difference between the diversity observed in molecular studies and the strains cultivated from natural systems is likely due to the loss of cell viability during sampling, or the enrichment and isolation protocols. The importance of using growth media compositions to target unique metabolic strategies and thus increase the diversity of cultured species has been discussed elsewhere (e.g., Alain and Querellou, 2009; Overmann et al., 2017). Furthermore, the idea of reducing sample exposure to conditions that cause cell death is regularly used, as is the case for obligate anaerobes, specifically methanogens. More recently, similar reasoning has been applied to the high-pressure biosphere; with novel technologies being deployed to capture and characterize microbes at elevated pressure (e.g., Garel et al., 2019; Peoples et al., 2019). Nonetheless, widespread application of high-pressure techniques throughout sampling, transfer, enrichment, and isolation is limited by challenges associated with cost, technology development, and technical expertise that are amplified when considering high-pressure sampling and cultivation (Cario et al., 2019). Furthermore, even when enrichments or isolates are cultivated at elevated pressures in laboratory settings, traditional methods that use static pressure vessels require decompression during subsampling (e.g., Takai et al., 2008), and the effects of decompression on experimental results for laboratory cultivation experiments has not been thoroughly or systematically investigated.
To explore the potential effects of decompression on microbial growth dynamics, we used two model sulfate reducers, D. salexigens and A. fulgidus to compare growth rates, cell yields, and maximum growth pressures between cultivation experiments that included several cycles of decompression and repressurization during subsampling, and those that maintained cultures at constant pressure throughout. Both strains showed significant exponential growth at elevated pressure for both types of cultivation conditions, although both had maximum growth rates at ambient pressure, classifying them as piezosensitive strains. We note that in previous studies we observed piezophilic behavior with maximum growth rates at 20 MPa (Oliver et al., 2020), and found that the growth rate measured at 0.1 MPa was particularly sensitive to the presence of a gas phase. Additionally, the maximum pressure for exponential growth for both strains was extended under isobaric conditions, confirming that cultivation techniques can impact experimentally determined growth characteristics of isolated strains.
In general, the impacts of the two cultivation techniques (isobaric vs. cyclic decompression) followed similar patterns for both D. salexigens and A. fulgidus, even though the specific pressure ranges, growth rates, and cell yields were strain-specific. Comparing cultivation techniques across both species, growth dynamics can be delineated into three different pressure regimes. The low-pressure regime (LP) has robust exponential growth with only marginal decreases in growth rates as pressures increase and little variation in maximum cell density. The high-pressure regime (HP) includes the range that exceeds the maximum pressure for exponential growth and is often characterized by cell death. The transitional pressure regime (TP) lies in between, is characterized by markedly lower growth rates and cell yields than those observed in the LP, and sometimes exponential growth is hard to discern. Within the TP the impacts of several cycles of decompression and repressurization are most noticeable for both growth rates and maximum cell yields.
The LP regime of D. salexigens is 0.1–20 and 0.1–40 MPa for A. fulgidus. For both strains, exponential growth rates decreased with increasing pressure, but maximum cell densities were similar at the end of the exponential growth phase. In this low-pressure regime, the effects of cyclic decompression are pronounced for growth rates, which are lower for cyclic decompression cultivation compares to isobaric experiments. However, little to no discernible impact was observed for cell densities. Growth rates and cell densities are more variable in the TP and HP regimes. For D. salexigens the transition pressure range is 30–40 MPa. While exponential growth is not discernible at 30 MPa for either condition, there are measurable increases in cell density late in the isobaric cultivation, while only cell death is observed with cyclic decompression cultivation (Figure 1A). It is also in the TP range that the largest difference in cell density is observed (Figure 1D). A similar pattern is observed for A. fulgidus for which the transition pressure range is 40–70 MPa. Exponential growth is observed in isobaric cultures up to 60 MPa, while the limit for cell division in cyclic decompression cultivation is 50 MPa. Overall, significantly slower growth rates (Figure 2C) were observed in A. fulgidus subjected to decompression from 40 to 60 MPa. Furthermore, the impact of decompression on cell density was also most apparent at 50 and 60 MPa (Figure 2D). Finally, while the HP regime is generally characterized by cell death (decreasing cell density with time) there are still noticeable differences between the two cultivation techniques. For D. salexigens no growth was obvious at 40 or 50 MPa, however cells subjected to cyclic decompression exhibited clearly higher rates of cell death (Figure 1A). Additionally for A. fulgidus, decreases in cell density with time were slower for isobaric cultivation.
The processes that might cause the largest differences in both growth rate and cell yield in the TP and HP are not immediately apparent. However, the robust exponential growth in the LP indicates that cells in this lower pressure range are not significantly impacted by these conditions. As both D. salexigens and A. fulgidus are piezosensitive strains that were isolated and optimized at ambient pressure conditions (0.1 MPa), it follows that decompression from moderate pressures to ambient pressure (their optimum pressure for growth) has little impact on growth rates and cell yields. However, for piezotolerant and piezophilic organisms, decompression would repeatedly expose cells to sub-optimal pressure conditions. In these cases, we hypothesize that cyclic decompression would have a more significant negative impact on growth. Additionally, if extended exposure to elevated pressure were inducing adaptation, even in a subpopulation of the culture, these cells would be selected against during decompression. Further characterization of these and other piezotolerant and piezophilic strains are critical next steps to better understand the effects of cyclic decompression on laboratory characterization of isolated species. Overall, we expect that the difference between isobaric cultivation and cultivation with cyclic decompression will be more pronounced in piezotolerant and piezophilic strains.
Concluding Remarks
For two deep biosphere species investigated here, A. fulgidus and D. salexigens, growth was negatively impacted by sample decompression, and these detriments to growth were more significant at pressures further from optimum pressures. These results highlight the need to maintain constant pressure without decompression during cultivation of high-pressure strains. Such an approach is likely to increase the pressure ranges of piezotolerant species, potentially reclassify other strains as piezophilic, and ultimately expand our knowledge of the diversity of known piezophiles, putting into perspective the habitat constraints for many strains whose pressure ranges have been underestimated. Even the piezophilic and piezotolerant organisms that are in culture have likely been isolated after decompression, inevitably selecting against strains more sensitive to decompression. For high-pressure samples that are decompressed and subsequently enriched at elevated in situ pressures, those strains that are somewhat tolerant to decompression, like A. fulgidus, would outcompete more sensitive strains, and results of any subsequent analyses would not necessarily be representative of the active in situ microbial communities.
Furthermore, maintaining in situ pressures becomes even more critical when retrieving samples from deep ecosystems, for example, 5–6 km—habitats more likely to host obligate piezophiles—for which decompression would subject native communities to pressure changes of over 50 MPa. Thus far, most of the obligate piezophiles identified have been psychrophilic bacteria sampled from depths greater that 6 km (i.e., ~60 MPa pressures; for example, Yayanos et al., 1981; Deming et al., 1988; Kato et al., 1998; Nogi et al., 2004) and only one obligate piezophilic hyperthermophilic archaeon, Pyrococcus yayanosii, has been identified from a hydrothermal vent at 4.1 km depth (Birrien et al., 2011). Given the extent of the subsurface biosphere, it is unlikely that these few obligate piezophiles represent the full diversity of Earth’s largest microbiome. Variable volume, high-pressure devices like the PUSH and others (reviewed in Cario et al., 2019), can be used not only to retrieve samples from up to ~9–10 km water depth, but all can be connected in series (as shown here and in Garel et al., 2019), so that sample transfer and enrichment can also be carried out without decompression. Such an approach will be more selective for piezophiles and obligate piezophiles, and greatly expand our understanding of the activity and physiology of the deep biosphere microbiome.
Data Availability Statement
The original contributions presented in the study are included in the article/Supplementary Material; further inquiries can be directed to the corresponding authors.
Author Contributions
GO, AC, and KR designed this research project and analyzed data. GO and AC performed the experiments and collected data and wrote the original draft. KR edited and wrote sections of the manuscript. All authors contributed to revisions of the manuscript, tables, and figures and approved the submitted version.
Funding
Funding for this work was provided by the NASA Exobiology and PSTAR Programs (NNX13AP2G9 and 80NSSC17K0252 to KR), the Deep Carbon Observatory (Subawards: 10371-07, 10561-01, and 10311-11 to KR), an NSF Graduate Fellowship (FAIN 1247271 and 1744655 to GO), and a GSA Research grant to GO. Additional support was provided by startup funds from Rensselaer Polytechnic Institute to KR.
Conflict of Interest
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
Publisher’s Note
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We thank Bruce Watson for generously gifting us four pressure vessels that allowed us to perform these growth experiments.
Supplementary Material
The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmicb.2022.867340/full#supplementary-material