INTRODUCTION

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LARGE VOLUMES OF GAS dissolve in the tissues of divers. The gases not metabolized by the diver, usually nitrogen or helium, are eliminated slowly during decompression. Gradual decompression lasting for days is needed following extended human dives to pressures of 20 atm or more, to avoid the excessive gas supersaturation and bubble formation that may lead to decompression sickness (DCS) (14). We describe a method for biochemically converting some of the dissolved gas to a nongaseous form. We call this method biochemical decompression.

Hydrogen has been used as the nonmetabolic portion of a breathing gas for deep hyperbaric exposures. Its lower density compared with helium reduces the effort of ventilation under high pressures (1, 2). Hydrogen is not metabolized by mammalian tissues (5, 12), but it can be readily converted to methane by microbes of the kingdom Archaea that can be found in the human colon (8). Methanobrevibacter smithii is the species responsible for the production of almost all human colonic methane; it uses H₂ to reduce carbon dioxide

(1)

Under normal circumstances, the H₂ is produced by colonic bacteria that ferment dietary plant polysaccharides (8). The magnitude of methane production in humans can be quite large. Some people produce up to three liters per day (15).

Utilizing the metabolism of M. smithii for our purposes is ideal: 4 mol of H₂ are converted to 2 mol of an innocuous, nongaseous molecule (water) and only 1 mol of a gas (methane) (Eq. 1). CO₂, the electron acceptor for this reaction, is abundant in mammalian tissues. Our approach was to place live suspensions of M. smithii into the proximal end of the large intestines of rats. While the rats breathed hyperbaric H₂, the metabolism of H₂ by the methanogen was demonstrated by monitoring the rate of methane release from the rats. With the use of a standardized compression and decompression sequence, efficacy at reducing DCS was confirmed by determining the incidence of DCS in rats with and without M. smithii treatments.

	METHODS

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Animals. Male Sprague-Dawley rats (Rattus norvegicus, n = 163, body mass range 215-324 g; Table 1) were used for all experiments. The rats were housed before experiments in an accredited animal care facility and had ad libitum access to rat chow (Prolab; Agway, Syracuse, NY) and water. All procedures were approved by an Animal Care and Use Committee. The experiments reported here were conducted according to the principles set forth in the Guide for the Care and Use of Laboratory Animals.

Analysis of variance was used to confirm that the various groups of control and treated animals did not differ from each other in body mass (P = 0.446). Maximum likelihood analysis was used to demonstrate that differences in body mass did not influence DCS outcome in these experiments.

Culturing of methanogen. M. smithii, strain PS, was obtained from the stock culture collection of the Wadsworth Center for Laboratories and Research, New York State Department of Health. This microbe was originally isolated from sewage. The microbe was cultured in 2-liter glass bottles modified with a serum bottle neck and side arm (9). The medium, which was based on rumen fluid (400 ml), was described elsewhere (10). Duplicate cultures were incubated at 37°C with stirring and 80% H₂-20% CO₂ at 1-2 atm. The gas within the culture bottles was replaced once daily. At 48 h, the cultures were centrifuged under H₂-CO₂ to concentrate the cells. The cells were resuspended in 12 ml O₂-free sodium bicarbonate solution (0.5 g NaHCO₃/100 ml) containing 0.005 M dithiothreitol (DTT) as a reducing agent. The suspension was stored under H₂-CO₂ at 4°C.

In vitro methanogenic activity was measured in duplicate with 0.2 ml of suspension and 0.3 ml of the bicarbonate-DTT solution in serum bottles with ~8 ml headspace volume. The bottles were pressurized to 2 atm with H₂-CO₂ and incubated on a platform shaker (280 excursions/min) for 1 h at 37°C. Methane and H₂ concentrations were determined by gas chromatography (11). The methane production rate per unit volume for the suspensions was retained in storage at 4°C for at least 8 days. In most experiments, the suspensions were used within 2 days of completion of culturing.

Surgical procedure to introduce methanogen. Sixty-three rats (Table 1) were anesthetized by inhalation of halothane (2-bromo-2-chloro-1,1,1-trifluoroethane). With the use of aseptic surgical techniques, an incision was made in the abdomen to expose the cecum. A cannula flushed with the bicarbonate-DTT solution was introduced from the rectum. The cannula was slowly passed through the entire length of the large intestine until it could be seen through the wall of the cecum. After delivering 2 ml of M. smithii suspension, the cannula was removed and the abdomen was closed with sutures and surgical clips. Torbugesic (0.04 mg/kg) was injected intramuscularly for postoperative pain. Animals were allowed to recover for 1-3 h.

Some animals treated with M. smithii (n = 34; Table 1) were used immediately after recovery in the experiments to measure methane. The remaining animals treated with M. smithii (n = 20; Table 1) were used immediately after recovery to test for DCS risk. Surgical control animals (n = 20 to test for methane, n = 20 to test for DCS risk; Table 1) underwent an identical procedure, except that the cecal injectate was 2 ml of 10% sodium bicarbonate.

Animals were removed from the study for only the following reasons: 1) when it was observed either during the surgery or over the course of the next 12-24 h that the cannulation procedure had damaged the colon (n = 6), 2) when the anaerobic conditions for the injection of microbes were not met due to improperly made DTT (n = 2), or 3) when animals experienced DCS immediately after their hyperbaric exposure for measuring methane (n = 10).

Dive protocol for measuring methane release rate. Four or five rats were placed together in a clear box (40-liter internal volume). Dry rat chow pellets and slices of apple were also put into the box to provide food and liquids. The box was placed inside a hyperbaric chamber (5.6 m³ internal volume, WSF Industries, Buffalo, NY) within a facility specially built for safe handling of high pressures of H₂ (following NFPA Code 50A for H₂ at consumer sites). Observation ports on the chamber allowed ad libitum viewing of the animals. A thermometer installed inside the rats' box was monitored throughout the dive. Chamber temperature was thermostatically controlled to keep the box at a temperature that appeared comfortable for the rats: ~28°C at 11 atm and 30°C at 23.7 atm.

The chamber was pressurized with He to an absolute pressure of 11 atm for 1 h. A stream of chamber gas (12-18 l/min STP, depending on gas composition) passed continuously through the animals' box and was sampled by a gas chromatograph (Hewlett-Packard 5890A, Series II, Wilmington, DE) every 12 min. This analysis of the ventilation of the box allowed regulation of chamber O₂, which could be added independently of other gases, to maintain normoxia (0.2-0.4 atm O₂) inside the box throughout the experiment. The He in the chamber was then replaced with H₂ at constant total pressure to a final concentration of 89.4 ± 0.6% H₂ (range 85.5-92.3%). This was accomplished primarily by opening chamber outlet and intake valves simultaneously and replacing the escaping chamber gases with H₂ and O₂ as needed to maintain constant pressure and normoxia. To speed the increase in H₂ concentration after concentration exceeded 50%, the chamber was pressurized two or three times with pure H₂ from 11 to 12 atm briefly and then returned to 11 atm. This exchange of H₂ for He was completed in ~2 h. The chamber was then further pressurized with H₂ to a total pressure of 23.7 atm (95 ± 0.6% H₂, range 90.7-98.1% H₂). This maximum pressure was maintained for 2.5 h. The chamber was then depressurized to 11 atm absolute pressure at 0.2-0.45 atm/min, which was intended to be slow enough to be unlikely to cause DCS in the rats (6). After 45 min at 11 atm, the H₂ in the chamber was replaced by He, maintaining a constant total pressure of 11 atm. In some cases, two or three excursions to 12 atm with pure He were used to speed the replacement of H₂ with He. To reduce the H₂ concentration to <4% typically required 2-2.5 h. The chamber was then further decompressed at 0.1-0.25 atm/min until the chamber had returned to 1 atm.

The animals were removed from the chamber, observed for overt signs of DCS, and left in cages with food and water overnight. Those animals with methanogen (n = 19; Table 1) or surgical controls (n = 18; Table 1) that appeared in normal health after a 12-h interval in 1-atm air were then tested for their risk of DCS, as described below.

Dive protocol for testing of decompression sickness risk. A separate dive protocol, which has been described in detail previously (3), was followed to test animals for their risk of DCS (Table 1). Five rats per test were placed inside a chamber (140-liter internal volume) that was pressurized to 10 atm with pure He at a rate of 1-2 atm/min. The chamber was flushed at constant pressure with a mixture of 2% O₂ in H₂. The chamber was further pressurized to 23.7 atm, and a final gas composition of 2% O₂, 4% He, 94% H₂. The rats remained at this pressure for 20 min, which was sufficient time to saturate the animals with H₂ (6). The animals were then decompressed within 36-40 s to a pressure of 10.8 atm and observed for signs of DCS over the next 30 min. While inside the chamber, rats were kept inside a drum mill made of wire mesh, rotating at 3.6 m/min, which obligated the rats to walk continuously throughout the experiment. The gait and position of rats on the mill was monitored through observation ports on the chamber. Signs of DCS were recorded as a limping or sideways gait, difficulty keeping up with the motion of the mill, falling, difficulty righting after falling, or rolling passively on the mill (3, 6). Two or three observers arrived at a consensus opinion to diagnose DCS, which required at least 1 min of signs.

All animals were euthanized in the chamber at 11 atm by introducing 2 atm CO₂. This terminated any discomfort for those rats with DCS.

	RESULTS

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Release of methane. Figure 1 presents the results of one experiment in which each of the five rats was supplied with a 2-ml suspension of M. smithii with an in vitro metabolic activity of 50 µmol H₂ uptake/min. The rats released an average of 0.76 ± 0.05 µmol CH₄/min per animal (mean ± SE 4 gas chromatographic measurements) during the initial hour at 11 atm, while their breathing mixture contained He rather than H₂ (Fig. 1). We presume that most of this methane was released as flatus, but any methane released with the breath or transcutaneously would have been sampled as well.

View larger version (26K): [in this window] [in a new window]   Fig. 1.   Average methane release rate per rat () from 5 rats with Methanobrevibacter smithii (50 µmol H2 uptake/min activity) in their large intestines as they breathed H2 at various pressures (). Points on abscissa are as follows. a: At 11 atm total pressure, before addition of H2, breathing a mixture of 88% helium in air. b: Commencement of flushing in H2 to replace the helium gas mixture at a constant total pressure of 11 atm. c: Increasing chamber pressure by addition of H2. d: Reaching 23.7 atm total pressure. e: Leaving 23.7 atm total pressure. f: Reaching 11 atm total pressure. g: Commencement of flushing in helium to replace the H2 gas mixture at a constant total pressure of 11 atm. Oxygen in the chamber was maintained at 0.2-0.4 atm O2 throughout.

Methane release rate increased, beginning within minutes of the appearance of H₂ in the chamber, reaching 2.21 ± 0.11 µmol CH₄/min per animal by the end of the H₂ exposure at 11 atm. As both chamber pressure and H₂ partial pressure increased, methane release rate increased again and reached 2.74 ± 0.12 µmol CH₄/min per animal by the end of the 2.5-h period at maximum pressure. Animals did not appear to be in any discomfort and often slept throughout large portions of the pressurization.

During chamber depressurization, methane release rate increased sharply, reaching a peak of 3.55 µmol CH₄/min per animal 45 min after depressurization was initiated (Fig. 1). Methane release rate then fell as both chamber pressure and H₂ partial pressure fell. There was no appearance of discomfort in the animals during depressurization; they were more wakeful and often groomed themselves.

This experimental procedure was executed a total of seven times, always using a 2-ml suspension of M. smithii per rat. In vitro activity for the suspensions ranged from 27 to 104 µmol H₂ uptake/min. Results from each procedure were qualitatively similar to those presented in Fig. 1, but there were quantitative differences in methane release rate that depended on the activity of methanogen delivered (Fig. 2). Animals supplied with doses of <50 µmol H₂ uptake/min activity (the dose for Fig. 1) released less methane throughout the experiment than illustrated in Fig. 1. However, animals supplied with >50 µmol H₂ uptake/min activity released approximately the same amounts of methane as shown in Fig. 1. Untreated control animals (n = 10; Table 1) and animals that had undergone the control surgical procedure (n = 20; Table 1) released no detectable methane while breathing either He or H₂ (limit of detection ~0.1 µmol CH₄/min).

View larger version (16K): [in this window] [in a new window]   Fig. 2.   Methane release rate from rats as a function of the in vitro activity of M. smithii placed in their intestines. Rats were breathing 91-98% H2 at 23.7 atm total pressure for 2.5 h. Each nonzero point represents the mean rate per animal for 5 animals during final hour of the H2 exposure. SEs of these mean values were 2-4% of the means (too small to be illustrated here). No detectable methane was released from untreated animals (n = 10) or from animals with control surgical procedures to place a sodium bicarbonate solution into their intestines (n = 20). Dotted line is illustrative only.

DCS risk. Animals supplied with M. smithii at a mean activity of 51 ± 16 µmol H₂ uptake/min (range 32-101 µmol H₂ uptake/min) and tested immediately for DCS risk had a 25% (5 of 20) incidence of DCS, brought on by the standardized sequence of compression to 23.7 atm and rapid decompression to 11 atm (Fig. 3A). This was significantly lower than the incidence of DCS for untreated rats of 56% (28 of 50; P < 0.01, ² one-sided test, Fig. 3A). Animals that underwent the control surgical procedure, followed immediately by the test for DCS risk, had a DCS incidence that was not significantly different from that of the untreated controls (65%; 13 of 20, P = 0.34, Fisher's exact test; Fig. 3A), but was significantly higher than that of the animals treated with methanogen (P < 0.01, ² one-sided test; Fig. 3A).

View larger version (22K): [in this window] [in a new window]   Fig. 3.   Incidence of decompression sickness for rats with and without M. smithii placed in their intestines, brought on by a standardized compression and decompression sequence. A: animals with no treatment (controls) and with a bicarbonate solution (surgical controls) or M. smithii (with methanogen) placed surgically in their intestines and tested within 1-3 h of surgery. B: animals with surgically implanted bicarbonate solution or M. smithii tested for DCS ~24 h after the surgery and 12 h after completion of an exposure to hyperbaric H2 to measure methane release rate. Error bars represent 95% binomial confidence limits.

The rats supplied with methanogen and tested first for their methane release rate and then for their risk of DCS (Table 1), ~24 h after the surgical delivery of M. smithii, had a DCS incidence of 36.8% (7 of 19; Fig. 3B). This incidence is not significantly different from that of animals tested within 1-3 h after treatment with M. smithii (Fig. 3A; P = 0.33; Fisher's exact test). However, it is significantly lower than that of untreated controls (Fig. 3A; P < 0.05, ² one-sided test). It is also significantly lower (P = 0.05, ² one-sided test) than the 55.6% (10 of 18; Fig. 3B) DCS incidence of animals subjected to the control surgical procedure, with an identical history of one pressurization to measure methane release rate and a second pressurization to measure DCS risk.

	DISCUSSION

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The release of detectable quantities of methane (Fig. 1) indicated that M. smithii had been delivered in a viable state by our surgical procedure. When the rats were breathing He, before any exposure to H₂ in their breathing mixture, the methanogen was evidently metabolizing H₂ released by endogenous H₂-producing bacteria (Ref. 8; Fig. 1). Although M. smithii is native to the gut flora of many mammals, it is not normally present in this strain of rats (7). This accounts for the absence of detectable methane release from untreated and surgical controls even while they were breathing H₂ (Fig. 2).

The increases in methane release rate with increasing H₂ partial pressure (Fig. 1) indicated that H₂ breathed by the rats was diffusing into the intestine and being metabolized by M. smithii. The increase in methane release rate was, however, not directly proportional to the increase in H₂ partial pressure; relatively more methane was produced per unit H₂ partial pressure at 11 atm than at 23.7 atm absolute pressure (Fig. 1). This suggests a limitation to H₂ delivery to the methanogen. Hydrogen delivery within the intestine is governed by diffusional processes through the intestinal wall and also by transport of dissolved H₂ to the intestine via the blood. Because diffusional processes are proportional to pressure gradients, this nonproportionality in methane release rate versus H₂ partial pressure suggests a perfusion limitation to H₂ delivery to the methanogen.

The sigmoidal relationship between in vitro activity of methanogen supplied to animals and methane release rate at maximum pressure (Fig. 2) also suggests that H₂ delivery to the microbe became rate limiting to their metabolism at the higher levels of methanogenic activity tested. Thus there is apparently a limit to the amount of H₂ that can be removed per unit time by a microbe in the intestine. If so, biochemical decompression cannot be extended indefinitely to greater levels of effectiveness by adding greater quantities or activities of microbes. A limitation of the magnitude found in these experiments may, however, be only an artifact of our experimental design. If the microbe were dispersed through a larger volume of the contents of the large intestine, or if drugs to increase intestinal blood flow were administered, it might be possible to increase the amount of H₂ eliminated per unit time by supplying higher methanogenic activity than used here.

We had not expected the spike in methane release shortly after initiating decompression (Fig. 1). This spike may reflect an increase in discharge of methane produced earlier in the dive and released by the rats as flatus either as a startle response to the depressurization of the chamber or as a simple phenomenon of the expansion of intestinal gases. However, in several of the 7 replications of this experiment, there was a second, smaller increase in methane release rate associated with the chamber H₂ washout, when there was a change in H₂ partial pressure but not total chamber pressure. We speculate that these periods of high methane release rate during decompression reflected bubble formation of H₂ in the cecum due to H₂ supersaturation. Gaseous H₂ may have supported a higher methanogenesis than when the H₂ supply was in physical solution in intestinal and surrounding tissue fluids during compression or constant chamber pressure.

The rate of methane release during decompression was ~20%-40% higher than the rate during compression at the same H₂ partial pressures (Fig. 1). This probably indicates that during compression there is methanogenesis primarily of H₂ recently inhaled by the rats, but during decompression there is additional methanogenesis of H₂ stored in the tissues of the animals, as well as H₂ recently inhaled. This observation is particularly encouraging: feasibility of biochemical decompression depends on reduction of tissue stores of inert gas before onset of DCS symptoms.

The treatment of rats with M. smithii was responsible for significantly lowering their risk of DCS from a chosen compression and decompression sequence to less than one-half of the risk in untreated controls (Fig. 3). The results from the surgical control groups showed that this decreased risk was not an artifact of the surgical procedure, either a few hours after surgery (Fig. 3A) or on the following day (Fig. 3B). The negative outcome of the surgical controls was critical because it has been documented that some activations of the immune system before a dive can reduce DCS risk (3). There appears to be a reduced inflammatory response to bubbles following a dive if a complement or some other element within the immune system is activated before the dive (for a review, see Ref. 3).

The protective effect against DCS of the treatments with M. smithii persisted for at least 24 h (Fig. 3B). However, M. smithii was probably not retained in a viable state for much longer than 24 h, due to loss of these alien microbes from the intestinal flora. Although fecal or cecal samples of some animals treated with M. smithii and left at 1 atm (Table 1) showed traces of methanogenic activity 24 h after the delivery of methanogen (7 of 8 animals sampled), no methanogenesis was observed after 2-8 days (0 of 3 animals sampled). It is significant that no health problems were observed that appeared to be caused by the methanogen itself. Animals kept at 1 atm after receiving methanogen treatment remained in good health and with feces normal in appearance during observation periods of days to weeks.

We are persuaded that there is a causal link between the methane released from rats treated with M. smithii (Figs. 1 and 2) and their reduced incidence of DCS (Fig. 3). As an approximation, assume that the average solubility of H₂ in the various tissues of a rat is 0.02 ml H₂ · ml tissues¹ · atm¹ (13). Assume also that H₂ production by bacteria in the gut flora is inhibited and falls to zero at elevated environmental H₂ pressures (8). A rat with a body mass of 250 g and a core body temperature of 38°C, breathing 91% H₂ at 23.7 atm total pressure (21.6 atm H₂ partial pressure) will contain 4.2 mmol H₂ when saturated. Saturation of a rat with H₂ is predicted to occur within 15 min (6). While the rats were breathing H₂ at this partial pressure, they released an average of 2.5 µmol CH₄/min when the in vitro activity of M. smithii placed in their intestines equaled or exceeded 50 µmol H₂ uptake/min (Fig. 2). (We use the average methane release rate after 2 h at maximum pressure, rather than the methane release rate in the first 20 min, because we do not know the lag time between methane production in the intestine and our sampling time.) Assuming that 4 µmol H₂ were consumed for each micromole of methane we detected (Eq. 1), we estimate that M. smithii was eliminating ~10 µmol H₂/min from the rats while at this pressure. Over a 20-min period, the methanogen thus eliminated ~200 µmol H₂. The H₂ elimination by the methanogen would thus reduce the total H₂ load in the rat by nearly 5%. This is equivalent to saturating a rat with H₂ at a 5% lower pressure than its actual ambient pressure; i.e., at 22.5 atm vs. 23.7 atm total pressure. A study in which DCS risk was computed as a function of pressure for rats breathing 2% O₂ and 4% He in H₂ and using a compression and decompression sequence identical to ours (6) estimated that the risk at 23.7 atm is ~60% and that at 22.5 atm is ~25%. These risk values correspond closely to those we actually measured for untreated rats versus those treated with M. smithii (Fig. 3).

This calculation utilizes several simplifying assumptions. In addition to those already listed, it ignores the volume of H₂ that diffuses from the environment into a subsaturated rat as a consequence of the increased H₂ partial pressure gradient generated by the methanogenic elimination of H₂ inside the rat. It also ignores the elevated rate of H₂ removal by the methanogen at the critical time during and immediately following decompression (Fig. 1). This simple calculation is nevertheless illustrative that the fluxes of H₂ under consideration are within the appropriate range to explain our results.

We envision that packaging M. smithii in an ingestible, enteric-coated capsule that targets delivery to the large intestine may make biochemical decompression applicable to human divers breathing H₂. Hydrogen diving is, however, restricted in its use to great depths and limited professional applications. The gas mixture most commonly used in sport, professional, and military diving is air. Thus even more appealing is the idea of replicating this approach using nitrogen-metabolizing bacteria (4). Biochemical decompression following hyperbaric exposures to air would revolutionize diving.

Perspectives