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       Hydrogen sulfide (H2S) has multiple physiological and pathological effects on the human body. Sodium hydrosulfide (NaHS) is widely used as a pharmacological tool to evaluate the effects of H2S in biological experiments. Although the loss of H2S from NaHS solutions takes only a few minutes, NaHS solutions have been used as donor compounds for H2S in drinking water in some animal studies. This study investigated whether drinking water with a NaHS concentration of 30 μM prepared in rat/mouse bottles could remain stable for at least 12–24 hours, as suggested by some authors. Prepare a solution of NaHS (30 μM) in drinking water and immediately pour it into rat/mouse water bottles. Samples were collected from the tip and the inside of the water bottle at 0, 1, 2, 3, 4, 5, 6, 12 and 24 hours to measure the sulfide content using the methylene blue method. In addition, male and female rats were injected with NaHS (30 μM) for two weeks and serum sulfide concentrations were measured every other day during the first week and at the end of the second week. The NaHS solution in the sample obtained from the tip of the water bottle was unstable; it decreased by 72% and 75% after 12 and 24 hours, respectively. In samples obtained from the inside of the water bottles, the decrease in NaHS was not significant within 2 hours; however, it decreased by 47% and 72% after 12 and 24 hours, respectively. NaHS injection did not affect the serum sulfide level of male and female rats. In conclusion, NaHS solutions prepared from drinking water should not be used for H2S donation because the solution is unstable. This route of administration will expose animals to irregular and smaller than expected amounts of NaHS.
       Hydrogen sulfide (H2S) has been used as a toxin since 1700; however, its possible role as an endogenous biosignaling molecule was described by Abe and Kimura in 1996. Over the past three decades, numerous functions of H2S in various human systems have been elucidated, leading to the realization that H2S donor molecules may have clinical applications in the treatment or management of certain diseases; see Chirino et al. for a recent review.
       Sodium hydrosulfide (NaHS) has been widely used as a pharmacological tool to assess the effects of H2S in many cell culture and animal studies5,6,7,8. However, NaHS is not an ideal H2S donor because it is rapidly converted to H2S/HS- in solution, is easily contaminated with polysulfides, and is easily oxidized and volatilized4,9. In many biological experiments, NaHS is dissolved in water, resulting in passive volatilization and loss of H2S10,11,12, spontaneous oxidation of H2S11,12,13, and photolysis14. Sulfide in the original solution is lost very rapidly due to volatilization of H2S11. In an open container, the half-life (t1/2) of H2S is about 5 minutes, and its concentration decreases by about 13% per minute10. Although the loss of hydrogen sulfide from NaHS solutions takes only a few minutes, some animal studies have used NaHS solutions as a source of hydrogen sulfide in drinking water for 1–21 weeks, replacing the NaHS-containing solution every 12–24 hours.15,16,17,18,19,20,21,22,23,24,25,26 This practice is not consistent with the principles of scientific research, since drug dosages should be based on their use in other species, especially humans.27
       Preclinical research in biomedicine aims to improve the quality of patient care or treatment outcomes. However, the results of most animal studies have not yet been translated to humans28,29,30. One of the reasons for this translational failure is the lack of attention to the methodological quality of animal studies30. Therefore, the aim of this study was to investigate whether 30 μM NaHS solutions prepared in rat/mouse water bottles could remain stable in drinking water for 12–24 h, as claimed or suggested in some studies.
       All experiments in this study were conducted in accordance with the published guidelines for the care and use of laboratory animals in Iran31. All experimental reports in this study also followed the ARRIVE guidelines32. The Ethics Committee of the Institute of Endocrine Sciences, Shahid Beheshti University of Medical Sciences, approved all experimental procedures in this study.
       Zinc acetate dihydrate (CAS: 5970-45-6) and anhydrous ferric chloride (CAS: 7705-08-0) were purchased from Biochem, Chemopahrama (Cosne-sur-Loire, France). Sodium hydrosulfide hydrate (CAS: 207683-19-0) and N,N-dimethyl-p-phenylenediamine (DMPD) (CAS: 535-47-0) were purchased from Sigma-Aldrich (St. Louis, MO, USA). Isoflurane was purchased from Piramal (Bethlehem, PA, USA). Hydrochloric acid (HCl) was purchased from Merck (Darmstadt, Germany).
       Prepare a solution of NaHS (30 μM) in drinking water and immediately pour it into the rat/mouse water bottles. This concentration was chosen based on numerous publications using NaHS as a source of H2S; see the Discussion section. NaHS is a hydrated molecule that can contain varying amounts of water of hydration (i.e., NaHS•xH2O); according to the manufacturer, the percentage of NaHS used in our study was 70.7% (i.e., NaHS•1.3 H2O), and we took this value into account in our calculations, where we used a molecular weight of 56.06 g/mol, which is the molecular weight of anhydrous NaHS. Water of hydration (also called water of crystallization) is the water molecules that make up the crystalline structure33. Hydrates have different physical and thermodynamic properties compared to anhydrates34.
       Before adding NaHS to the drinking water, measure the pH and temperature of the solvent. Immediately pour the NaHS solution into the rat/mouse water bottle in the animal cage. Samples were collected from the tip and from the inside of the water bottle at 0, 1, 2, 3, 4, 5, 6, 12, and 24 h to measure sulfide content. Sulfide measurements were taken immediately after each sampling. We obtained samples from the tip of the tube because some studies have shown that the small pore size of the water tube can minimize H2S evaporation15,19. This issue appears to apply to the solution in the bottle as well. However, this was not the case for the solution in the neck of the water bottle, which had a higher evaporation rate and was autoxidizing; in fact, the animals drank this water first.
       Male and female Wistar rats were used in the study. The rats were housed in polypropylene cages (2–3 rats per cage) under standard conditions (temperature 21–26 °C, humidity 32–40%) with 12 h of light (7 am to 7 pm) and 12 h of darkness (7 pm to 7 am). The rats had free access to tap water and were fed with standard chow (Khorak Dam Pars Company, Tehran, Iran). Age-matched (6 months) female (n=10, body weight: 190–230 g) and male (n=10, body weight: 320–370 g) Wistar rats were randomly divided into control and NaHS (30 μM) treated groups (n=5 per group). To determine the sample size, we used the KISS (Keep It Simple, Stupid) approach, which combines previous experience and power analysis35. We first conducted a pilot study on 3 rats and determined the mean serum total sulfide level and standard deviation (8.1 ± 0.81 μM). Then, considering 80% power and assuming a two-sided 5% significance level, we determined a preliminary sample size (n = 5 based on previous literature) that corresponded to a standardized effect size of 2.02 with the predefined value suggested by Festing for calculating the sample size of experimental animals35. After multiplying this value by the SD (2.02 × 0.81), the predicted detectable effect size (1.6 μM) was 20%, which is acceptable. This means that n = 5/group is sufficient to detect a 20% mean change between groups. Rats were randomly divided into control and NaSH-treated groups using the random function of Excel software 36 (Supplementary Fig. 1). Blinding was performed at the outcome level, and the investigators performing the biochemical measurements were not aware of the group assignments.
       The NaHS groups of both sexes were treated with 30 μM NaHS solution prepared in drinking water for 2 weeks; Fresh solution was supplied every 24 h, during which time body weight was measured. Blood samples were collected from the tail tips of all rats under isoflurane anesthesia every other day at the end of the first and second weeks. Blood samples were centrifuged at 3000 g for 10 min, serum was separated and stored at –80°C for subsequent measurement of serum urea, creatinine (Cr), and total sulfide. Serum urea was determined by enzymatic urease method, and serum creatinine was determined by photometric Jaffe method using commercially available kits (Man Company, Tehran, Iran) and an automatic analyzer (Selectra E, serial number 0-2124, The Netherlands). The intra- and interassay coefficients of variation for urea and Cr were less than 2.5%.
       The methylene blue (MB) method is used to measure total sulfide in drinking water and serum containing NaHS; MB is the most commonly used method for measuring sulfide in bulk solutions and biological samples11,37. The MB method can be used to estimate the total sulfide pool38 and measure inorganic sulfides in the form of H2S, HS- and S2 in the aqueous phase39. In this method, sulfur is precipitated as zinc sulfide (ZnS) in the presence of zinc acetate11,38. Zinc acetate precipitation is the most widely used method for separating sulfides from other chromophores11. ZnS was redissolved using HCl11 under strongly acidic conditions. The sulfide reacts with DMPD in a stoichiometric ratio of 1:2 in a reaction catalyzed by ferric chloride (Fe3+ acts as an oxidizing agent) to form the dye MB, which is detected spectrophotometrically at 670 nm40,41. The detection limit of the MB method is approximately 1 μM11.
       In this study, 100 μL of each sample (solution or serum) was added to a tube; then 200 μL of zinc acetate (1% w/v in distilled water), 100 μL of DMPD (20 mM in 7.2 M HCl), and 133 μL of FeCl3 (30 mM in 1.2 M HCl) were added. The mixture was incubated at 37°C in the dark for 30 min. The solution was centrifuged at 10,000 g for 10 min, and the absorbance of the supernatant was read at 670 nm using a microplate reader (BioTek, MQX2000R2, Winooski, VT, USA). Sulfide concentrations were determined using a calibration curve of NaHS (0–100 μM) in ddH2O (Supplementary Fig. 2). All solutions used for the measurements were freshly prepared. The intra- and interassay coefficients of variation for sulfide measurements were 2.8% and 3.4%, respectively. We also determined the total sulfide recovered from sodium thiosulfate-containing drinking water and serum samples using the fortified sample method42. The recoveries for sodium thiosulfate-containing drinking water and serum samples were 91 ± 1.1% (n = 6) and 93 ± 2.4% (n = 6), respectively.
       Statistical analysis was performed using GraphPad Prism software version 8.0.2 for Windows (GraphPad Software, San Diego, CA, USA, www.graphpad.com). A paired t-test was used to compare the temperature and pH of drinking water before and after NaHS addition. The loss of H2S in the NaHS-containing solution was calculated as a percentage decrease from baseline uptake, and to assess whether the loss was statistically significant, we performed a one-way repeated-measures ANOVA followed by Dunnett’s multiple comparison test. Body weight, serum urea, serum creatinine, and total serum sulfide over time were compared between control and NaHS-treated rats of different sexes using a two-way mixed (between-within) ANOVA followed by a Bonferroni post hoc test. Two-tailed P values ​​< 0.05 were considered statistically significant.
       The pH of drinking water was 7.60 ± 0.01 before NaHS addition and 7.71 ± 0.03 after NaHS addition (n = 13, p = 0.0029). The temperature of drinking water was 26.5 ± 0.2 and decreased to 26.2 ± 0.2 after NaHS addition (n = 13, p = 0.0128). Prepare 30 μM NaHS solution in drinking water and store it in a water bottle. NaHS solution is unstable and its concentration decreases over time. When sampling from the neck of the water bottle, a significant decrease (68.0%) was observed within the first hour, and the NaHS content in the solution decreased by 72% and 75% after 12 and 24 hours, respectively. In samples obtained from water bottles, the reduction in NaHS was not significant up to 2 hours, but after 12 and 24 hours it had decreased by 47% and 72%, respectively. These data indicate that the percentage of NaHS in a 30 μM solution prepared in drinking water had decreased to approximately one-quarter of the initial value after 24 hours, regardless of sampling location (Figure 1).
       Stability of NaHS solution (30 μM) in drinking water in rat/mouse bottles. After solution preparation, samples were taken from the tip and the interior of the water bottle. Data are presented as mean ± SD (n = 6/group). * and #, P < 0.05 compared with time 0. The photograph of the water bottle shows the tip (with opening) and the body of the bottle. The tip volume is approximately 740 μL.
       The concentration of NaHS in the freshly prepared 30 μM solution was 30.3 ± 0.4 μM (range: 28.7–31.9 μM, n = 12). However, after 24 h, the concentration of NaHS decreased to a lower value (mean: 3.0 ± 0.6 μM). As shown in Figure 2, the concentrations of NaHS to which the rats were exposed were not constant during the study period.
       The body weight of female rats increased significantly over time (from 205.2 ± 5.2 g to 213.8 ± 7.0 g in the control group and from 204.0 ± 8.6 g to 211.8 ± 7.5 g in the NaHS-treated group); however, NaHS treatment had no effect on body weight (Fig. 3). The body weight of male rats increased significantly over time (from 338.6 ± 8.3 g to 352.4 ± 6.0 g in the control group and from 352.4 ± 5.9 g to 363.2 ± 4.3 g in the NaHS-treated group); however, NaHS treatment had no effect on body weight (Fig. 3).
       Changes in body weight in female and male rats after administration of NaHS (30 μM). Data are presented as mean ± SEM and were compared using two-way mixed (within-between) analysis of variance with Bonferroni post hoc test. n = 5 of each sex in each group.
       Serum urea and creatine phosphate concentrations were comparable in control and NaSH-treated rats throughout the study. Furthermore, NaSH treatment did not affect serum urea and creatinechrome concentrations (Table 1).
       Baseline serum total sulfide concentrations were comparable between control and NaHS-treated male (8.1 ± 0.5 μM vs. 9.3 ± 0.2 μM) and female (9.1 ± 1.0 μM vs. 6.1 ± 1.1 μM) rats. NaHS administration for 14 days had no effect on serum total sulfide levels in either male or female rats (Fig. 4).
       Changes in serum total sulfide concentrations in male and female rats after administration of NaHS (30 μM). Data are presented as mean ± SEM and were compared using a two-way mixed (within-within) analysis of variance with Bonferroni post hoc test. Each sex, n = 5/group.
       The main conclusion of this study is that drinking water containing NaHS is unstable: only about a quarter of the initial total sulfide content can be detected 24 hours after sampling from the tip and inside of rat/mouse water bottles. Furthermore, rats were exposed to unstable NaHS concentrations due to the loss of H2S in the NaHS solution, and the addition of NaHS to drinking water did not affect body weight, serum urea and creatine chromium, or total serum sulfide.
       In this study, the rate of H2S loss from 30 μM NaHS solutions prepared in drinking water was approximately 3% per hour. In a buffered solution (100 μM sodium sulfide in 10 mM PBS, pH 7.4), the sulfide concentration was reported to decrease by 7% over time over 8 h11. We have previously defended intraperitoneal administration of NaHS by reporting that the rate of sulfide loss from a 54 μM NaHS solution in drinking water was approximately 2.3% per hour (4%/hour in the first 12 h and 1.4%/hour in the last 12 h after preparation)8. Earlier studies43 found a constant loss of H2S from NaHS solutions, primarily due to volatilization and oxidation. Even without the addition of bubbles, sulfide in the stock solution is rapidly lost due to H2S volatilization11. Studies have shown that during the dilution process, which takes about 30–60 seconds, about 5–10% of H2S is lost due to evaporation6. To prevent evaporation of H2S from the solution, researchers have taken several measures, including gentle stirring of the solution12, covering the stock solution with plastic film6, and minimizing exposure of the solution to air, since the rate of H2S evaporation depends on the air-liquid interface.13 Spontaneous oxidation of H2S occurs mainly due to transition metal ions, especially ferric iron, which are impurities in water.13 Oxidation of H2S results in the formation of polysulfides (sulfur atoms linked by covalent bonds)11. To avoid its oxidation, solutions containing H2S are prepared in deoxygenated solvents44,45 and then purged with argon or nitrogen for 20–30 min to ensure deoxygenation.11,12,37,44,45,46 Diethylenetriaminepentaacetic acid (DTPA) is a metal chelator (10–4 M) that prevents HS- autoxidation in aerobic solutions. In the absence of DTPA, the autoxidation rate of HS- is approximately 50% over approximately 3 h at 25°C37,47. Furthermore, since the oxidation of 1e-sulfide is catalyzed by ultraviolet light, the solution should be stored on ice and protected from light11.
       As shown in Figure 5, NaHS dissociates into Na+ and HS-6 when dissolved in water; this dissociation is determined by the pK1 of the reaction, which is temperature dependent: pK1 = 3.122 + 1132/T, where T ranges from 5 to 30°C and is measured in degrees Kelvin (K), K = °C + 273.1548. HS- has a high pK2 (pK2 = 19), so at pH < 96.49, S2- is not formed or is formed in very small amounts. In contrast, HS- acts as a base and accepts H+ from an H2O molecule, and H2O acts as an acid and is converted to H2S and OH-.
       Formation of dissolved H2S gas in NaHS solution (30 µM). aq, aqueous solution; g, gas; l, liquid. All calculations assume that water pH = 7.0 and water temperature = 20 °C. Created with BioRender.com.
       Despite evidence that NaHS solutions are unstable, several animal studies have used NaHS solutions in drinking water as an H2S donor compound15,16,17,18,19,20,21,22,23,24,25,26 with intervention durations ranging from 1 to 21 weeks (Table 2). During these studies, the NaHS solution was renewed every 12 h, 15, 17, 18, 24, 25 h or 24 h, 19, 20, 21, 22, 23 h. Our results showed that rats were exposed to unstable drug concentrations due to the loss of H2S from the NaHS solution, and the NaHS content in rats’ drinking water fluctuated significantly over 12 or 24 h (see Figure 2). Two of these studies reported that H2S levels in water remained stable over 24 h22 or that only 2–3% H2S losses were observed over 12 h15, but they did not provide supporting data or measurement details. Two studies have shown that the small diameter of water bottles can minimize H2S evaporation15,19. However, our results showed that this may only delay H2S loss from a water bottle by 2 h rather than 12–24 h. Both studies note that we assume that the NaHS level in the drinking water did not change because we did not observe a color change in the water; therefore, oxidation of H2S by air was not significant19,20. Surprisingly, this subjective method assesses the stability of NaHS in water rather than measuring the change in its concentration over time.
       The loss of H2S in NaHS solution is related to pH and temperature. As noted in our study, dissolving NaHS in water results in the formation of an alkaline solution50. When NaHS is dissolved in water, the formation of dissolved H2S gas depends on the pH value6. The lower the pH of the solution, the greater the proportion of NaHS present as H2S gas molecules and the more sulfide is lost from the aqueous solution11. None of these studies reported the pH of drinking water used as a solvent for NaHS. According to WHO recommendations, which are adopted by most countries, the pH of drinking water should be in the range 6.5–8.551. In this pH range, the rate of spontaneous oxidation of H2S increases approximately tenfold13. Dissolving NaHS in water in this pH range will result in a dissolved H2S gas concentration of 1 to 22.5 μM, which emphasizes the importance of monitoring the pH of the water before dissolving NaHS. In addition, the temperature range reported in the above study (18–26 °C) would result in a change in the concentration of dissolved H2S gas in the solution of approximately 10%, since temperature changes alter pK1, and small changes in pK1 can have a significant impact on the concentration of dissolved H2S gas48. In addition, the long duration of some studies (5 months)22, during which large temperature variability is expected, also exacerbates this problem.
       All studies except one21 used 30 μM NaHS solution in drinking water. To explain the dose used (i.e. 30 μM), some authors pointed out that NaHS in the aqueous phase produces exactly the same concentration of H2S gas and that the physiological range of H2S is 10 to 100 μM, so this dose is within the physiological range15,16. Others explained that 30 μM NaHS can maintain the plasma H2S level within the physiological range, i.e. 5–300 μM19,20. We consider the concentration of NaHS in water of 30 μM (pH = 7.0, T = 20 °C), which was used in some studies to study the effects of H2S. We can calculate that the concentration of dissolved H2S gas is 14.7 μM, which is about 50% of the initial NaHS concentration. This value is similar to the value calculated by other authors under the same conditions13,48.
       In our study, NaHS administration did not change the body weight; this result is consistent with the results of other studies in male mice22,23 and male rats18; However, two studies reported that NaSH restored the decreased body weight in nephrectomized rats24,26, whereas other studies did not report the effect of NaSH administration on body weight15,16,17,19,20,21,25. Furthermore, in our study, NaSH administration did not affect the serum urea and creatine chromium levels, which is consistent with the results of another report25.
       The study found that addition of NaHS to drinking water for 2 weeks did not affect total serum sulfide concentrations in male and female rats. This finding is consistent with the results of Sen et al. (16): 8 weeks of treatment with 30 μM NaHS in drinking water did not affect plasma sulfide levels in control rats; however, they reported that this intervention restored the decreased H2S levels in plasma of nephrectomized mice. Li et al. (22) also reported that treatment with 30 μM NaHS in drinking water for 5 months increased plasma free sulfide levels in aged mice by about 26%. Other studies have not reported changes in circulating sulfide after addition of NaHS to drinking water.
       Seven studies reported using Sigma NaHS15,16,19,20,21,22,23 but did not provide further details on the water of hydration, and five studies did not mention the source of NaHS used in their preparation methods17,18,24,25,26. NaHS is a hydrated molecule and its water of hydration content can vary, which affects the amount of NaHS required to prepare a solution of a given molarity. For example, the NaHS content in our study was NaHS•1.3 H2O. Thus, the actual NaHS concentrations in these studies may be lower than those reported.
       “How can such a short-lived compound have such a long-lasting effect?” Pozgay et al.21 asked this question when evaluating the effects of NaHS on colitis in mice. They hope that future studies will be able to answer this question and speculate that NaHS solutions may contain more stable polysulfides in addition to the H2S and disulfides that mediate the effect of NaHS21. Another possibility is that very low concentrations of NaHS remaining in solution may also have a beneficial effect. In fact, Olson et al. provided evidence that micromolar levels of H2S in the blood are not physiological and should be in the nanomolar range or absent altogether13. H2S may act through protein sulfation, a reversible post-translational modification that affects the function, stability, and localization of many proteins52,53,54. In fact, under physiological conditions, approximately 10% to 25% of many liver proteins are sulfylated53. Both studies acknowledge the rapid destruction of NaHS19,23 but surprisingly state that “we controlled the concentration of NaHS in drinking water by replacing it daily.”23 One study accidentally stated that “NaHS is a standard H2S donor and is commonly used in clinical practice to replace H2S itself.”18
       The above discussion shows that NaHS is lost from solution through volatilization, oxidation and photolysis, and therefore some suggestions are made to reduce the loss of H2S from solution. First, the evaporation of H2S depends on the gas-liquid interface13 and the pH of the solution11; therefore, to minimize the evaporative loss, the neck of the water bottle can be made as small as possible as described previously15,19, and the pH of the water can be adjusted to an acceptable upper limit (i.e., 6.5–8.551) to minimize the evaporative loss11. Second, spontaneous oxidation of H2S occurs due to the effects of oxygen and the presence of transition metal ions in drinking water13, so deoxygenation of drinking water with argon or nitrogen44,45 and the use of metal chelators37,47 can reduce the oxidation of sulfides. Third, to prevent the photodecomposition of H2S, water bottles can be wrapped with aluminum foil; This practice also applies to light-sensitive materials such as streptozotocin55. Finally, inorganic sulfide salts (NaHS, Na2S, and CaS) can be administered via gavage rather than dissolved in drinking water as previously reported56,57,58; studies have shown that radioactive sodium sulfide administered via gavage to rats is well absorbed and distributed to virtually all tissues59. To date, most studies have administered inorganic sulfide salts intraperitoneally; however, this route is rarely used in clinical settings60. On the other hand, the oral route is the most common and preferred route of administration in humans61. Therefore, we recommend evaluating the effects of H2S donors in rodents by oral gavage.
       A limitation is that we measured sulfide in aqueous solution and serum using the MB method. The methods for measuring sulfide include iodine titration, spectrophotometry, electrochemical method (potentiometry, amperometry, coulometric method and amperometric method) and chromatography (gas chromatography and high-performance liquid chromatography), among which the most commonly used method is the MB spectrophotometric method62. A limitation of the MB method for measuring H2S in biological samples is that it measures all sulfur-containing compounds and not free H2S63 because it is performed under acidic conditions, which results in the extraction of sulfur from the biological source64. However, according to the American Public Health Association, MB is the standard method for measuring sulfide in water65. Therefore, this limitation does not affect our main results on the instability of solutions containing NaHS. Furthermore, in our study, the recovery of sulfide measurements in water and serum samples containing NaHS was 91% and 93%, respectively. These values ​​are in line with previously reported ranges (77–92)66, indicating acceptable analytical precision42. It is worth noting that we used both male and female rats in accordance with National Institutes of Health (NIH) guidelines to avoid overreliance on male-only animal studies in preclinical studies67 and to include both male and female rats whenever possible68. This point has been emphasized by others69,70,71.
       In conclusion, the results of this study indicate that NaHS solutions prepared from drinking water cannot be used to generate H2S due to their instability. This route of administration would expose animals to unstable and lower than expected levels of NaHS; therefore, the findings may not be applicable to humans.
       The datasets used and/or analyzed during the current study are available from the corresponding author upon reasonable request.
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