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and illustrations in this method, please contact the SLTC at (801) 233-4900.
These procedures were designed and tested for internal use by OSHA personnel. Mention of any company name or commercial product does not constitute endorsement by OSHA.
HYDROGEN SULFIDE IN WORKPLACE ATMOSPHERES
|OSHA Permissible Exposure Limits
Hydrogen Sulfide (Final Rule Limits):
10 ppm (Time Weighted Average)15 ppm (Short-Term Exposure Limit)
| Hydrogen Sulfide
20 ppm (Ceiling)50 ppm (Peak - 10 Min Exposure)
||A calibrated personal sampling pump is used to draw air through a filter impregnated with silver nitrate which converts the hydrogen sulfide to silver sulfide.
|Recommended Air Volume:
||2 to 6 L
|Recommended Sampling Rate
Peak, Ceiling, or STEL Samples:
Time Weighted Average Samples:
0.2 L/min0.1 L/min
||The sample is analyzed as sulfide by differential pulse polarography.
0.40 ppm (2-L air volume)0.90 ppm (2-L air volume)
|Precision and Accuracy
Validation Level: CVT Bias Overall Error
10.1 to 39.2 ppm0.038-0.031±10.7%
|Date (Date Revised):
||1983 (December, 1989)
Commercial manufacturers and products mentioned in this method are for
descriptive use only and do not constitute endorsements by USDOL-OSHA.
Similar products from other sources can be substituted.
Branch of Inorganic Methods Development
OSHA Technical Center
Salt Lake City, Utah
This method describes the collection of airborne hydrogen sulfide (H2S) in the workplace on a
silver nitrate (AgNO3)-impregnated filter and analysis by differential pulse polarography (DPP).
2. Range and Detection Limit (8.6)
Previously, H2S was collected in a midget impinger containing an alkaline suspension of cadmium
hydroxide. The sulfide was precipitated as cadmium sulfide (CdS) and subsequently analyzed by the methylene blue
calorimetric procedure (8.1). Due to the photosensitivity of CdS, it was necessary to
protect the impinger sample from light at all times. Also, the impinger base and stem could contain CdS deposits after
sample collection. This required the entire impinger sample be sent to the lab for analysis. Problems associated with
impinger sampling were additional incentives to develop a more acceptable sampling method.
Hydrogen sulfide is collected on a Whatman 4 filter paper (Whatman Labsales, Hillsboro, OR) which has been impregnated with
AgNO3. The H2S reacts with the AgNO3 to form
silver sulfide, a greyish-black precipitate (8.2,
silver sulfide is dissolved in an alkaline cyanide solution and analyzed for sulfide by DPP using a dropping mercury
1.3. Advantages and Disadvantages
1.3.1. The sampling device is small, portable, and involves no liquids. Humidity does not significantly affect the sampling efficiency of the device.
1.4. Physical and chemical properties (8.4,
1.3.2. The sulfide formed is stable and non-volatile. Desorption and preparation of samples for analysis involve simple procedures.
1.3.3. Collected samples are analyzed by means of a quick analytical method.
1.3.4. This method has adequate sensitivity for measuring workplace atmospheric concentrations of
H2S for Short-Term Exposure Limit (STEL), Ceiling, or Time Weighted Average (TWA) measurements.
1.3.5. The analysis is specific for sulfide in the presence of other organic or inorganic sulfur compounds.
1.3.6. One disadvantage of the method is the amount of H2S collected is limited by the capacity
of the AgNO3-impregnated filter.
1.3.7. Another disadvantage is the necessity of protecting the impregnated filters from light at all times. The
AgNO3-impregnated filters darken upon exposure to light.
1.3.8. Alkaline cyanide solutions are used in the sample preparation and analytical procedure. Safety precautions must be
followed during their use and disposal.
1.3.9. The alkaline cyanide solutions may contain some background sulfide which must be subtracted from the amount
of sulfide found in each sample. The precision of the method is affected by the reproducibility of the background sulfide.
Hydrogen sulfide (CAS No. 7783-06-4) is a colorless, poisonous, flammable gas with a characteristic odor of rotten eggs at
low concentrations. Collapse, coma, and death from respiratory failure may result from brief exposure at high concentrations.
Exposure at low concentrations produces irritation of conjunctiva and mucous membranes. It is soluble in water, alcohol, ether,
petroleum solvents, and crude petroleum. Some physical properties are listed:
||1.192 (air = 1)
|Explosive range in air
||4.5 - 45.5%
|Olfactory fatigue level
1.5. Occupations with Potential Exposure to Hydrogen Sulfide (8.4)
|Animal fat and oil processors
|Animal manure removers
|Artificial flavor makers
|Asphalt storage workers
|Barium carbonate makers
||Manhole and trench workers
|Blast furnace workers
||Natural gas production and
|| processing workers
||Painters using polsulfide
|Carbon disulfide makers
|| caulking compounds
|Chemical laboratory workers,
||Petroleum production and
| teachers, students
|| refinery workers
|Citrus root fumigators
|Coal gasification workers
||Pipeline maintenance workers
|Coke oven workers
||Septic tank cleaners
||Sewage treatment plant workers
|Fermentation process workers
|Fishing and fish-processing workers
| and production workers
||Sugar beet and cane processors
||Sulfur spa workers
||Sulfur products processors
|Hydrochloric acid purifiers
|Hydrogen sulfide production
| and sales workers
|Lead ore sulfidizers
||Well diggers and cleaners
2.1. The analytical working range is from 0.05 to 4 µg/mL as sulfide.
3. Method Performance (8.6)
2.2. The qualitative and quantitative detection limits of the analytical method for a 50 mL sample extraction volume are
1.0 and 2.5 µg H2S, respectively. These values correspond to 0.4 ppm and 0.90 ppm
H2S when using a 2 L air volume.
3.1. This method was evaluated at high (85 to 88%) RH over the range of 10.1 to 39.2 ppm at an approximate temperature and
atmospheric pressure of 25.5°C and 640 mmHg, respectively. Samples were taken for 10 min using a 0.2 L/min sampling
3.2. The pooled coefficient of variation (CVT) for the overall sampling and analytical method in
the range tested was 0.038 and bias was -0.031.
3.3. In validation experiments, this method was capable of measurements within ±25% of the true value at least 95% of
the time. The overall error of the method was +10.7%.
3.4. The collection efficiency at high (85-88%) RH was 100%. A concentration of 40 ppm was used. Breakthrough tests were
conducted at both high (86%) and low (18%) RH at the same concentration of 40 ppm. At either RH, 6% breakthrough of H2S occurred after 20 min at a sampling rate of 0.2 L/min.
3.5. In storage stability studies, the mean recovery of samples analyzed after 30 days was within 3% of the mean recovery
of similar samples analyzed immediately after collection. Sample concentrations were about 20 ppm. The samples were stored
at normal lab temperatures in a dark environment.
4.1. The presence of other inorganic sulfide compounds in the atmosphere will interfere with the analysis of
4.2. Any substance in the atmosphere which oxidizes the sulfide formed on the impregnated filter can be a negative
interference. Literature studies suggest the presence of gases such as SO2,
CO2, 02, NO2, and
NH3 in the atmosphere will not affect the recovery of H2S (
4.3. Any compound with the same peak potential as sulfide when using the analytical conditions described in this method is
an interference. Studies suggest that organic and other inorganic sulfur compounds (sulfites, thiosulfates, mercaptans,
etc.) will not interfere in the analysis (8.7,
4.4. If an interference exists, changing the operating conditions of the polarograph or the electrolyte may circumvent the problem.
4.5. When other substances are known or suspected to be present in the air sampled, the identities of the substances should be transmitted with the sample.
5.1.1. Sample assembly:
5.2. Sampling Procedure
Filter holder consisting of a two-piece cassette, 37-mm diameter.
Backup pad, 37-mm, cellulose.
Silver nitrate-impregnated cellulose filters, 37-mm (see the Appendix if preparation is necessary)
5.1.2. Gel bands (Omega Specialty Instrument Co., Chelmsford, MA) for sealing cassettes.
5.1.3. Sampling pumps capable of sampling at 0.1 to 0.2 liters per minute (L/min).
5.1.4. Assorted flexible tubing.
5.1.5. Stopwatch and bubble tube or meter for pump calibration.
(Note: The impregnated filters should be handled with nonmetallic forceps. Also, the AgNO3-impregnated filters are light-sensitive and can turn black upon prolonged exposure to light.)
5.2.1. Place an impregnated filter and a cellulose backup pad into a 37 mm polystyrene two-piece cassette filter holder. Seal the cassette with a shrinkable gel band. Place plastic end caps on the cassette. Shield the cassette from light by completely wrapping it with black tape or aluminum foil. Store the cassette in a dark environment until use.
5.2.2. Calibrate each personal sampling pump with a prepared cassette in-line at the flow rate listed below:
|Type of Sample
|Ceiling, STEL, or Peak
5.2.3. Attach prepared cassettes to calibrated sampling pumps (the backup pad should face the pump) and place in
appropriate positions on the employee or workplace area. Begin sampling; due to potential breakthrough,
do not exceed the recommended sampling times listed above.
5.2.4. Place plastic end caps on each cassette after sampling.
5.2.5. If possible, refrigerate the collected samples during storage periods prior to analysis. Samples do not need to be
refrigerated during handling or shipping.
Submit at least one blank sample with each set of air samples. Blank filter samples should be handled in the same manner
as other samples, except no air is drawn through the blank. Attach an OSHA-21 seal around each cassette in such a way as
to secure the end caps. Send the samples to the laboratory with the OSHA 91A paperwork requesting hydrogen sulfide analysis.
6.1. Safety Precautions
6.1.1. Safety glasses, labcoat, and gloves must be worn at all times.
6.1.2. Cyanide compounds and solutions are powerful poisons which prevent the utilization of oxygen by the body tissues.
Care must be exercised when using these compounds. They can be absorbed through the skin. Cyanides which contact the skin
should be washed off immediately.
6.1.3. Mercury vapor and liquid are very toxic substances which can be readily absorbed through the respiratory tract and
the skin. Avoid skin contact with mercury and clean up mercury spills immediately.
6.1.4. The analyst must properly dispose of all reagents, standards, prepared samples, and waste solutions after their use.
Disposal of solutions containing no mercury is accomplished by pouring the solution down a sink with copious amounts
of tap water and rinsing the containing vessel several times with DI H2O. Waste solutions of
mercury and cyanide from the polarograph cell cups should be flushed in a large glass beaker with copious amounts of DI
H2O in a sink before placing the waste mercury in a metal container specifically designed for
Exercise care to prevent any mercury from entering the sink. Rinse the empty used polarograph cell cups several times with
DI H2O before cleaning. Never acidify the solutions or dispose in drains which are used for
acid disposal. Any acidic environment has the potential to convert the cyanide or sulfide salts to deadly hydrogen
cyanide and/or H2S gas.
6.1.5. Waste solutions containing CdS precipitate should be poured into a labeled plastic waste bottle containing 0.1 M
NaOH. The waste bottle should be properly disposed of. Check state and federal regulations for proper disposal. Never pour
this waste solution down the sink.
6.1.6. Refer to the Standard Operating Procedure (SOP) (8.9) and instrument manufacturer
manuals for proper operation of the polarographic instrument and safety precautions.
6.2.1. Polarographic Analyzer or Controller, Model 384 or 384B, (Princeton Applied Research (PAR), Princeton, NJ) with a
Model 303 or 303A dropping mercury electrode.
6.3. Reagents: All chemicals should be reagent grade or better.
6.2.2. Reference salt bridge tube (PAR Model K0154) filled with a saturated potassium chloride solution.
6.2.3. Saturated calomel reference electrode (PAR Model K0077) for use with the reference electrode salt bridge tube.
6.2.4. Glass or polypropylene polarographic cells, 15-mL.
6.2.5. Nitrogen purification system: Gas purifier for deoxygenating nitrogen, (Oxiclear, part no. DGP-250, Labclear, Oakland, CA. As an alternative, an oxygen scrubber can be constructed using a vanadous chloride solution as described in reference
6.2.6. Sulfide specific ion electrode (Model 94-16A, Orion Research, Inc. Cambridge, MA).
6.2.7. Double junction reference electrode (Model 90-02, Orion Research) with inner and outer filling solutions (Cat. Nos.
90-00-02 and 90-00-03, respectively).
6.2.8. Millivolt meter.
6.2.9. Magnetic stirrer and Teflon stirring bars.
6.2.10. Burette, glass, 10-mL.
6.2.11. Beakers, polypropylene, 100-mL, with tight fitting covers.
6.2.12. Bottle, narrow mouth, amber, linear polyethylene, opaque, 125- or 250-mL, with screw cap - used for storage of light-sensitive material.
6.2.13. Bottles, wide mouth, polypropylene, with screwcaps.
6.2.14. Nonmetallic forceps.
6.2.15. Glass rod.
6.2.16. Polystyrene disposable beakers.
6.2.17. Automatic pipette, adjustable, 20- to 5000-µL range (Gilson Pipetman, Rainin Instruments Inc.).
6.2.18. Volumetric pipettes, volumetric flasks, beakers and other laboratory glassware.
6.2.19. Tissue paper.
6.2.20. Analytical balance (0.01 mg).
6.3.1. Deionized water (DI H2O): Deionized, filtered, deoxygenated water is needed for
preparation of all solutions which will be used in the analysis. It is recommended to purge the DI
H2O with purified nitrogen before use.
6.4. Standard Preparation
6.3.2. Sodium hydroxide (NaOH).
6.3.3. Sodium hydroxide, 0.1 M solution. Dissolve 4.0 g of N in approximately 400 mL of DI H2O
and then dilute the solution to 1 L. Store the solution in a wide-mouth polypropylene bottle.
6.3.4. Sodium cyanide (NaCN), containing < 0.001% sulfide as an impurity.
6.3.5. Alkaline 0.5 M NaCN solution. Dissolve 2.0 g of NaOH and then 12.25 g of NaCN in approximately 300 mL of DI
H2O and then dilute the solution to 500 mL. Immediately and carefully transfer the solution to a wide-mouth, polypropylene bottle.
6.3.6. Alkaline 0.25 M NaCN solution. Dilute 250 ml of the 0.5 M NaCN solution to 500 mL with 0.1 M NaOH solution.
Immediately and carefully transfer the solution to a wide-mouth polypropylene bottle.
6.3.7. Nitric acid (HNO3), concentrated.
6.3.8. Nitric acid, 6 M: Carefully dilute 384 mL of concentrated HNO3 to 1 L using DI
6.3.9. Nitric acid, 1 M: Dilute 6.4 mL of concentrated HNO3 to 100 mL with DI
6.3.10. Sodium sulfide nonahydrate (Na2S·9H2O), crystalline.
6.3.11. Sodium sulfide (Na2S) stock solution (equivalent to 1,000 µg/mL
1. Weigh 0.7050 g of Na2S·9H2O crystals in a 50 mL disposable
polystyrene beaker (Note: Before weighing, remove any surface impurities by rinsing the crystals with DI H2O in an exhaust hood.
Quickly blot the crystals dry with tissue paper. Handle the crystals with nonmetallic forceps.).
6.3.12. Cadmium sulfate (CdSO4) anhydrous.
2. Dissolve the weighed Na2S·9H2O crystals in a disposable beaker containing about 30 mL of 0.1 M NaOH.
3. Quantitatively transfer the solution to a 100 mL volumetric flask. Rinse the beaker several times with 0.1 M NaOH and transfer to the volumetric
flask. Dilute the flask to the mark with 0.1 M NaOH.
4. Wrap the flask with aluminum foil to protect it from light. Refrigerate and keep the flask in a dark environment prior to and immediately after each use.
5. Prepare this solution weekly.
6.3.13. Cadmium sulfate (CdSO4), 0.1500 M standard solution (used in the standardization of the Na2S stock solution).
1. Heat approximately 3.5 g of anhydrous CdSO4 at 110°C for 24 h in a drying oven. Cool the dried CdSO4 in a desiccator for approximately 30 min.
6.3.14. Saturated potassium chloride solution for filling the salt bridge.
2. Quickly weigh 3.127 g of the dried CdSO4 in a 50 mL disposable polystyrene beaker and then dissolve in approximately 25 mL of DI H2O.
3. Quantitatively transfer this solution to a 100 mL volumetric flask. Rinse the beaker several times with DI H2O and transfer the rinses to the volumetric flask. Dilute to volume with DI H20.
6.4.1. To standardize the Na2S stock solution a titration is performed with a standard 0.1500 M CdSO4 solution. A sulfide ISE is used as the indicator. An orange-colored precipitate (CdS) is formed from the reaction of Cd2+ with S2- (8.11).
6.5. Sample Preparation
1. Connect the sulfide ion specific and double junction reference electrodes to the millivolt meter.
Place the electrodes in a 0.1 M NaOH solution to reduce stabilization time.
2. Allow the Na2S stock solution to come to ambient temperature and then pipette 25 mL of this
solution into a 50 mL disposable beaker containing a Teflon stirring bar. Use magnetic stirring throughout the titration.
3. Place the electrodes in the beaker and record the initial mV reading. Place a piece of insulating material such as
cardboard or Styrofoam between the magnetic stirrer and the beaker to minimize the heating of the standard solution by the stirrer.
4. Record the electrode potential as a function of titrant volume added. Initially add the 0.1500 m CdSO4 standard
titration solution from a 10 mL burette in 0.5 to 1.0 mL increments to the beaker. When the potential change per increment begins to
increase, add 0.05 mL to 0.2 ml of the titrant to the beaker. Add about 1 mL of the titrant beyond the endpoint.
5. The end point is the point of greatest slope on the titration curve. This can be determined by plotting the electrode
potential versus titrant volume or tabulating the change in potential per mL increment of titrant added.
6. Calculate the µg/mL equivalent of H2S in the Na2S stock solution:
|H2S (µg/mL) =
||A × B × 32.064 × 1,000 × 1.0629
||molarity of the standard CdSO4 solution.
||mL of standard CdSO4 solution required to titrate the Na2S stock solution.
||milligrams per milliequivalent sulfide.
||micrograms per milligram
||gravimetric factor (H2S/S2-)
||25.0 mL = mL of the Na2S solution used.
6.4.2. Prepare standards equivalent to 100 and 10 µg/mL H2S by making appropriate serial dilutions of the Na2S stock solution with the 0.1 M NaOH solution. Protect these standards from light and prepare daily.
Wash all glassware and plasticware, excluding the polarograph cell cups, in detergent, rinse in dilute (1 M) nitric acid, rinse thoroughly with DI H2O, and then air dry prior to use. Wash the cell cups in 6 M HNO3 and rinse with DI H2O before use.
6.5.1. Open the filter cassette. Carefully remove the sample filter with nonmetallic forceps and place it in a 100 mL polypropylene beaker.
6.6. Analytical Procedure
6.5.2. Immediately pipette 25 mL of the alkaline 0.5 M NaCN solution into the beaker and then tightly cover it with a polypropylene lid.
6.5.3. Place the covered beaker in a dark environment at ambient room temperature for approximately 6 h. Occasionally swirl the sample solution with a gentle motion. The silver sulfide on the filter will dissolve in the cyanide solution during this time.
6.5.4. Store the covered sample beaker overnight in a refrigerator.
6.5.5. Remove the beaker from the refrigerator and place in a dark environment for approximately 45 min to allow the sample solution to equilibrate to room temperature.
6.5.6. Pipette 25 mL of the 0.1 M NaOH solution into the sample beaker and then stir the sample solution with a clean glass rod. Cover each sample solution with a plastic lid until analysis. The solutions should be analyzed sometime during the same work day.
6.6.1. Protect and electrically isolate the silver reference electrode wire on the DME by installing a clean, empty reference electrode glass sleeve with a Vycor tip.
6.7. Analytical Recommendations
6.6.2. Install the filled salt bridge tube on the electrode support block. Insert the saturated calomel reference electrode (SCE) into the salt bridge tube and then connect the SCE cable to the reference electrode input.
6.6.3. Refer to reference
8.9 for operating the polarograph.
6.6.4. Turn on the nitrogen to a pressure of about 2 psi.
6.6.5. Initially set the following values (Note: Some of the settings mentioned below are instrument specific. Refer to specific operating and service manuals for other types of polarographs):
6.6.6. The differential pulse polarogram of sulfide using a DME yields a peak at approximately -0.690 V. The peak potential depends on reference electrode response and the DME operating parameters.
6.6.7. Prepare standards in the range of 0.05 to 4.0 µg/mL immediately before analysis by diluting µL aliquots of the Na2S working standards to 10 ml in the polarograph cell cups with the alkaline 0.25 M NaCN solution. A suggested method of preparation of the standard solutions in the cell cups is given:
|SS H2S Concn*
||Aliquot of SS
||0.25 M NaCN
||Final H2S Concn
|* All working standards are prepared in 0.25 M NaCN/0-1 N NaOH solution.
** Reagent Blank contains 9.8 mL NaCN/0.1 N NaOH and 0.2 mL of 0.1 N NaOH.
SS = Na2S stock solution. All H2S concns listed are as equivalent concns.
6.6.8. Scan a 1 µg/mL standard to determine the actual sulfide peak potential. If the peak is significantly different from -0.69 V, reset the peak, initial, and final potentials to window the peak (±0.080 V from peak).
6.6.9. Observe in this polarogram the rapidly rising baseline when at a more positive potential than the sulfide peak. This rising baseline is due to cyanide in the supporting electrolyte which has a peak potential of approximately -0.36 V. This baseline is minimized by a reagent blank subtraction routine found in the polarograph software.
6.6.10. Pipette 10 mL aliquots of the 50 mL sample solutions from the polypropylene beakers into the polarograph cell cups immediately prior to analysis.
6.6.11. Record the peak potential (V) and the peak current (µA) for each standard and sample.
6.7.1. Verify that mercury is flowing properly through the glass capillary of the DME before the analysis begins and during the analysis by observing mercury drop formation.
6.7.2. Because the sensitivity and linearity range increase with increasing cyanide concentration, it is important the cyanide concentration in the standard and analytical solutions be identical.
6.7.3. Analyze the reagent blank, a series of standards in the range of interest, and the samples. A freshly prepared standard should be analyzed after every five or six samples to monitor polarograph performance. Always rinse the electrodes with DI H2O and blot dry with a clean dry tissue before analyzing the next solution.
7.1. Determine the µg/mL H2S content of each sample and blank using a concentration-response linear regression curve (µA vs. µg/mL).
7.2. Calculate the total micrograms contained in each sample:
µg H2S = [(Sample Vol, mL) × µg/mL H2S]
7.3. Each air sample is blank corrected (µg sample - µg blank) and the concentration is then calculated to determine H2S exposure using the following equation:
7.4. Reporting Results
|ppm H2S =
||MV × µg H2Sb
molecular weight × air volume, L
|MV (Molar Volume)
||24.45 (25°C and 760 mmHg)
||Blank corrected sample result
|Molecular Weight (H2S)
This equation reduces to:
|ppm H2S =
||0.717 × µg sample
air volume, L
Results are reported to the industrial hygienist as ppm H2S.
8.1 National Institute for Occupational Safety and Health: NIOSH Manual of Analytical Methods. 2nd ed., Vol. 2 (DHEW/NIOSH Pub. No. 77-157-B). Cincinnati, OH: National Institute for Occupational Safety and Health, 1977. pp. S4-1-S4-10.
8.2 Natusch, D.F.S., H.B. Klonis, H.D. Axelrod, R.D. Teck and J.P. Lodge, Jr.: Sensitive Method for Measurement of Atmospheric Hydrogen Sulfide. Anal. Chem. 44: 2067-2070 (1972).
8.3 Natusch, D.F.S., J.R. Sewell and R.L. Tanner: Determination of Hydrogen Sulfide in Air -- An Assessment of Impregnated Paper Tape Methods. Anal. Chem. 46: 410-415 (1974).
8.4 National Institute for Occupational Safety and Health: Criteria for a Recommended Standard -- Occupational Exposure to Hydrogen Sulfide (DHEW/NIOSH Pub. No. 77-158). Cincinnati, OH: National Institute for Occupational Safety and Health, 1977.
8.5 Fassett, D.W. and D.D. Irish, ed.: Patty's Industrial Hygiene and Toxicology. 2nd rev. ed., Vol. 2. New York: John Wiley and Sons, 1963.
8.6 Occupational Safety and Health Administration Tedmical Center: Hydrogen Sulfide Backup Data Report (ID-141). Salt Lake City, UT. Revised 1989.
8.7 Renard, J.J., G. Kubes and H.I. Bolker: Polarographic Determination of Sulfur Compounds in Pulping Liquors. Anal. Chem. 47: 1347-1352 (1975).
8.8 Noel, D.L.: Sulfur Compounds in Kraft Pulping Liquor. Tappi 61: 73-76 (1978).
8.9 Occupational Safety and Health Administration Technical Center: Standard Operating Procedure for Polarography. Salt Lake City, UT. In progress (unpublished).
8.10 Princeton Applied Research: Application note 108, Why Dearation... and How. Princeton, NJ: Princeton Applied Research, 1974.
8.11 Ehman, D.L.: Determination of Parts-per-Billion Levels of Hydrogen Sulfide in Air by Potentiometric Titration with a Sulfide Ion-Selective Electrode as an Indicator. Anal. Chem. 48: 918-920 (1976).
Preparation of H2S Sampling Devices - Impregnated Filters
Preparation of filter impregnation solution, 2% AgNO3
Reagents (All reagents should be reagent grade or better):
Silver nitrate (AgNO3).
- Dissolve 2 g AgNO3, in approximately 60 mL DI H2O contained in a glass beaker. Immediately transfer this solution to a 100-mL volumetric flask.
- Pipette 1 mL of 1 N HNO3, 5 mL of glycerin, and 20 mL of ethanol into the 100-mL volumetric flask. Dilute the solution to volume with DI H2O.
- Immediately transfer the AgNO3 solution to an amber polyethylene bottle. Store the bottle in a dark environment. The AgNo3 solution has at least a 6 month shelf-life.
Preparation of Impregnated Filters
Whatman No. 4 (Special Order, Whatman Labsales, Hillsboro, OR) cellulose filters (37-mm diameter) are impregnated with the AgNO3 solution described above. This filter is prepared as follows:
- Place the cellulose filter over one of the holes of a polypropylene test tube rack or scintillation vial support rack. The rack should have hole diameters of 30-mm.
- Using subdued light, pipette 250 µL of the AgNO3 impregnation solution on the filter.
- Carefully transfer the filters and support to a dark environment and allow to dry at ambient temperature for approximately 120 min.
- Handle the filters in subdued light.
- Store the filters together in a dark environment at normal lab temperatures in tightly covered polystyrene petri dishes. Filters can be stored for at least 3 months.
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