Analytic Methods for the Oil and Gas 

Extraction Point Source Category

U.S. Environmental Protection Agency

Engineering and Analysis Division

Office of Water

1200 Pennsylvania Avenue, NW

Washington, D.C. 20460

December 2011

EPA-821-R-11-004
CONTENTS (Continued)

CONTENTS

Page

1. SUMMARY OF METHODS ...................................................................................................1-1

1.1 Static Sheen Test (EPA Method 1617)................................................................1-5

1.2 Drilling Fluids Toxicity Test (EPA Method 1619)..............................................1-5

1.3 Procedure for Mixing Base Fluids with Sediments (EPA Method 1646)............1-5

1.4 Protocol for the Determination of Degradation of Non-Aqueous Base 

Fluids in a Marine Closed Bottle Biodegradation Test System: Modified 

ISO 11734:1995 (EPA Method 1647) .................................................................1-6

1.5 Determination of Crude Oil Contamination in Non-Aqueous Drilling 

Fluids by Gas Chromatography/Mass Spectrometry (GC/MS) (EPA 

Method 1655).......................................................................................................1-7

1.6 Reverse Phase Extraction (RPE) Method for Detection of Oil 

Contamination in Non-Aqueous Drilling Fluids (NAF) (EPA Method 

1670) ....................................................................................................................1-7

1.7 Determination of the Amount Of Non-Aqueous Drilling Fluid (NAF) Base 

Fluid from Drill Cuttings by a Retort Chamber (Derived From API 

Recommended Practice 13B–2) (EPA Method 1674).........................................1-8

1.8 PAH Content of Oil by HPLC/UV (EPA Method 1654, Revision A).................1-8

1.9 Sediment Toxicity Test for NAF and SBM (EPA Method 1644) .......................1-8

1.10 Previous Publication of Oil and Gas Extraction Point Source Category

Analytic Methods.................................................................................................1-9

2. STATIC SHEEN TEST (EPA METHOD 1617).....................................................................2-11

2.1 Scope and Application .......................................................................................2-11

2.2 Summary of Method ..........................................................................................2-11

2.3 Interferences.......................................................................................................2-11

2.4 Apparatus, Materials, and Reagents...................................................................2-11

2.5 Calibration..........................................................................................................2-12

2.6 Quality Control Procedures................................................................................2-12

2.7 Sample Collection and Handling .......................................................................2-12

2.8 Procedure ...........................................................................................................2-13

3. DRILLING FLUIDS TOXICITY TEST (EPA METHOD 1619)..................................................3-1

3.1 Scope and Application .........................................................................................3-1

3.2 Summary of Method ............................................................................................3-1

3.3 Sample Collection................................................................................................3-1

3.4 Suspended Particulate Phase Sample Preparation ...............................................3-2

3.5 Guidance for Performing Suspended Particulate Phase Toxicity Tests 

Using Mysidopsis bahia.......................................................................................3-4

3.6 Methods for Positive Control Tests (Reference Toxicant)..................................3-7

3.7 Randomization Procedure....................................................................................3-7

3.8 References..........................................................................................................3-12

4. PROCEDURE FOR MIXING BASE FLUIDS WITH SEDIMENTS (EPA METHOD 1646) ............4-1

4.1 Determining the Wet to Dry Ratio for the Control Sediment..............................4-1

4.2 Determining the Density of the Wet Control or Dilution Sediment....................4-1

4.3 Determining the Amount of Base Fluid Needed..................................................4-1

4.4 Primary Mixing....................................................................................................4-2
4.5 Testing for Homogeneity of Base Fluid...............................................................4-2

4.6 Commencing the Sediment Toxicity Test............................................................4-2

4.7 References............................................................................................................4-3

5. PROTOCOL FOR THE DETERMINATION OF DEGRADATION OF NON-AQUEOUS BASE 

FLUIDS IN A MARINE CLOSED BOTTLE BIODEGRADATION TEST SYSTEM: MODIFIED 

ISO 11734:1995 (EPA METHOD 1647) ............................................................................5-1

5.1 Summary of Method ............................................................................................5-1

5.2 System Requirements...........................................................................................5-1

5.3 Test Set Up...........................................................................................................5-2

5.4 Concentration Verification Chemical Analyses...................................................5-6

5.5 Gas Monitoring Procedures .................................................................................5-7

5.6 Test Acceptability and Interpretation...................................................................5-9

5.7 Methane Measurement.........................................................................................5-9

5.8 Concentration Verification Analysis..................................................................5-11

5.9 Program Quality Assurance and Quality Control ..............................................5-12

6. DETERMINATION OF CRUDE OIL CONTAMINATION IN NON-AQUEOUS DRILLING 

FLUIDS BY GAS CHROMATOGRAPHY/MASS SPECTROMETRY (GC/MS) (EPA

METHOD 1655).................................................................................................................6-1

6.1 Scope and Application .........................................................................................6-1

6.2 Summary of Method ............................................................................................6-1

6.3 Definitions............................................................................................................6-2

6.4 Interferences and Limitations ..............................................................................6-2

6.5 Safety ...................................................................................................................6-2

6.6 Apparatus and Materials......................................................................................6-3

6.7 Reagents and Standards .......................................................................................6-4

6.8 Sample Collection Preservation and Storage.......................................................6-6

6.9 Quality Control ....................................................................................................6-6

6.10 Calibration............................................................................................................6-9

6.11 Procedure ...........................................................................................................6-11

6.12 Calculations........................................................................................................6-15

6.13 Method Performance..........................................................................................6-16

6.14 Pollution Prevention...........................................................................................6-16

6.15 Waste Management............................................................................................6-16

6.16 References..........................................................................................................6-17

6.17 Schematic Flowchart for Qualitative Identification...........................................6-18

7. REVERSE PHASE EXTRACTION (RPE) METHOD FOR DETECTION OF OIL 

CONTAMINATION IN NON-AQUEOUS DRILLING FLUIDS (NAF) (EPA METHOD 1670)......7-1

7.1 Scope and Application .........................................................................................7-1

7.2 Summary of Method ............................................................................................7-1

7.3 Definitions............................................................................................................7-1

7.4 Interferences.........................................................................................................7-2

7.5 Safety ...................................................................................................................7-2

7.6 Equipment and Supplies ......................................................................................7-2

7.7 Reagents and Standards .......................................................................................7-4

7.8 Sample Collection, Preservation, and Storage.....................................................7-4

7.9 Quality Control ....................................................................................................7-4
7.10 Calibration and Standardization...........................................................................7-7

7.11 Procedure .............................................................................................................7-7

7.12 Data Analysis and Calculations...........................................................................7-9

7.13 Method Performance............................................................................................7-9

7.14 Pollution Prevention.............................................................................................7-9

7.15 Waste Management............................................................................................7-10

7.16 References..........................................................................................................7-10

8. DETERMINATION OF THE AMOUNT OF NON-AQUEOUS DRILLING FLUID (NAF) BASE 

FLUID FROM DRILL CUTTINGS BY A RETORT CHAMBER (DERIVED FROM API

RECOMMENDED PRACTICE 13B–2) (EPA METHOD 1674)................................................8-1

8.1 Description...........................................................................................................8-1

8.2 Equipment............................................................................................................8-1

8.3 Procedure .............................................................................................................8-2

8.4 Calculations..........................................................................................................8-3

8.5 Requirements for Sampling Cuttings Discharge Streams for use with this 

Method .................................................................................................................8-9

8.6 Best Management Practices (BMPs) for use with this Method .........................8-12

9. PAH CONTENT OF OIL BY HPLC/UV (EPA METHOD 1654, REVISION A).......................9-1

9.1 Scope and Application .........................................................................................9-1

9.2 Summary of Method ............................................................................................9-1

9.3 Interferences.........................................................................................................9-1

9.4 Safety ...................................................................................................................9-2

9.5 Apparatus and Materials......................................................................................9-2

9.6 Reagents...............................................................................................................9-4

9.7 Calibration............................................................................................................9-5

9.8 Quality Assurance/Quality Control......................................................................9-6

9.9 Sample Collection, Preservation, and Handling ..................................................9-9

9.10 Dilution of Oil and Extracts.................................................................................9-9

9.11 High-Performance Liquid Chromatography......................................................9-10

9.12 HPLC System and Laboratory Performance......................................................9-11

9.13 Qualitative Identification ...................................................................................9-12

9.14 Quantitative Determination................................................................................9-12

9.15 Method Performance..........................................................................................9-13

9.16 References..........................................................................................................9-13

10. METHOD FOR CONDUCTING A SEDIMENT TOXICITY TEST WITH LEPTOCHEIRUS 

PLUMULOSUS AND NON-AQUEOUS DRILLING FLUIDS OR SYNTHETIC-BASED 

DRILLING MUDS (EPA METHOD 1644) ..........................................................................10-1

10.1 Summary of Method ..........................................................................................10-1

10.2 Test Requirements and Materials.......................................................................10-1

10.3 Procedures..........................................................................................................10-6

10.4 Recommended Test Conditions.........................................................................10-8

10.5 Biological Data ................................................................................................10-10

10.6 Quality Control ................................................................................................10-10

10.7 Reference Tests................................................................................................10-11

10.8 Interpretation of Result ....................................................................................10-12

10.9 References........................................................................................................10-13
v

LIST OF TABLES

Page

Table 1-1. Methods by Waste and Pollutant, Subpart A − Offshore Subcategory...................... 1-2

Table 1-2. Methods by Waste and Pollutant, Subpart D − Coastal Subcategory ........................ 1-4

Table 1-3. EPA Method Numbers for Oil and Gas Extraction Point Source Category

Analytical Methods and Prior CFR References in 40 CFR 435 ............................. 1-10

Table 3-1. Example of a Randomization Schedule...................................................................... 3-9

Table 3-2. Listing of Acute Toxicity Test Data (August 1983 to September 1983) With 

Eight Generic Drilling Fluids and Mysid Shrimp................................................... 3-11

Table 3-3. Partial Toxicity Test Passing Criteria....................................................................... 3-12

Table 5-1. Test Acceptability Criteria ......................................................................................... 5-9

Table 6-1. Gas Chromatograph/Mass Spectrometer (GC/MS) Operation Conditions................ 6-9

Table 6-2. Approximate Retention Time for Compounds......................................................... 6-10

Table 6-3. Recommended Ion Mass Numbers........................................................................... 6-13

Table 9-1. Performance Data and Method Acceptance Criteria for PAH ................................. 9-14

Table 9-2. HPLC Calibration Data ............................................................................................ 9-14

Table 10-1. Culture conditions for L. plumulosus. Conditions listed are consistent with 

culture conditions specified in ASTM E 1367-92 and subsequent updates (E 

1367-99).................................................................................................................. 10-3

Table 10-2. Dry food portion of the diet that has been used to successfully culture L. 

plumulosus .............................................................................................................. 10-4

Table 10-3. Conditions for conducting 96-hour NAF and 10-day SBM sediment toxicity

tests with L. plumulosus. Conditions listed are consistent with test conditions 

specified in ASTM E 1367- 92 and subsequent updates (E 1367-99) unless 

otherwise noted ....................................................................................................... 10-9

Table 10-4. Test acceptability requirements for 10-day NAF and 96-hr SBM tests with L. 

plumulosus. Requirements listed are consistent with those specified in ASTM 

E 1367-92 and subsequent updates (E 1367-99)................................................... 10-11
vi

LIST OF FIGURES

Page

Figure 3-1. Mysid Randomization Procedure.............................................................................. 3-8

Figure 6-1. Schematic Flowchart for Qualitative Identification............................................... 6-18

Figure 9-1. Liquid Chromatography of the Three-Component Standard and of No. 2 

Diesel Oil ................................................................................................................ 9-15
1-1

1. SUMMARY OF METHODS 

This document is a compilation of all but one of the analytical methods necessary for 

demonstrating compliance with the requirements of 40 CFR Part 435, effluent limitations 

guidelines and standards for the Oil and Gas Extraction Point Source Category. These methods 

are tests for regulated pollutant parameters. The waste sources and pollutant parameters 

regulated in 40 CFR Part 435 are listed in the following tables along with the required analytical 

or test method, and the subpart that cites the method. Table 1-1 presents the waste sources and 

pollutant parameters regulated by Subpart A – Offshore Subcategory. Table 1-2 presents the 

same information for Subpart D – Coastal Subcategory.

These methods were developed during two rulemakings: Offshore Subcategory (March 4, 1993; 

58 FR 12454) and the Synthetic-based Drilling Fluids (SBF) Effluent Guidelines (January 22, 

2001; 66 FR 6849). These analytic methods were developed in consultation and collaboration 

with industry. For example, the analytic methods for the Offshore Subcategory were developed 

with industry in the 1980’s and included the Static Sheen Test and Drilling Fluids Toxicity Test

(see August 26, 1985; 50 FR 34596). EPA again worked very closely with industry and adopted 

their recommendations in promulgating additional analytical methods for the Synthetic-based 

Drilling Fluids Effluent Guidelines (see January 22, 2001; 66 FR 6893). These additional 

analytical methods included tests for PAH content (as phenanthrene), sediment toxicity, 

biodegradation rate, and percentage of base fluid on drill cuttings (API retort method). EPA 

listed the analytic methods and in some cases re-printed the analytic methods in the Code of 

Federal Regulations (CFR). 

As previously noted there is one method that is not included in this document, which relates to 

Cook Inlet, Alaska, operators that are regulated by the Coastal subcategory (Subpart D). The 

Coastal subcategory prohibits the discharge non-aqueous drill cuttings unless there are technical 

limitations. Appendix 1 of 40 CFR Part 435 Subpart D provides the method for permit writers to 

determine when operators qualify for an exemption from the discharge prohibition. For those 

operators that quality for the exemption from the discharge prohibition EPA requires that these 

operators meet the same stock limitations and discharge limitations for drill cuttings associated 

with non-aqueous drilling fluids for operators in Offshore waters (see 40 CFR §435.13 and 

§435.15) in order to discharge drill cuttings associated with non-aqueous drilling fluids. This 

method remains in 40 CFR 435 and is not included in this document. 

The remainder of this section provides a summary of the analytical methods used for Part 435. 
1-2

Table 1-1. Methods by Waste and Pollutant, Subpart A − Offshore Subcategory

Waste source

Regulated Pollutant 

Parameter Analytical/Test Method

Cited in 40 CFR 

Part

Produced Water Oil & grease 40 CFR Part 136 §435.12, §435.13, 

§435.14, §435.15 

Water-based drilling 

fluids and associated 

drill cuttings

SPP Toxicity Drilling Fluids Toxicity Test 

EPA Method 1619

§435.13, §435.15

Free oil Static Sheen Test

EPA Method 1617

§435.12, §435.13, 

§435.14, §435.15

Diesel oil † ASTM specification D975-91 §435.13, §435.15

Mercury 40 CFR Part 136 §435.13, §435.15

Cadmium 40 CFR Part 136 §435.13, §435.15

Drill cuttings 

associated with nonaqueous drilling 

fluids: Discharge 

limitations

Diesel oil † ASTM specification D975-91 §435.13, §435.15

Free oil Static Sheen Test

EPA Method 1617

§435.12, §435.14

SPP Toxicity Drilling Fluids Toxicity Test 

EPA Method 1619

§435.13, §435.15

Drilling fluid sediment 

toxicity (4-Day Test)

Method for Conducting a Sediment Toxicity 

Test with Leptocheirus Plumulosus and NonAqueous Drilling Fluids or Synthetic-Based 

Drilling Muds (EPA Method 1644)

after using

Procedure for Mixing Base Fluids with 

Sediments

EPA Method 1646

§435.13, §435.15

Formation oil Reverse Phase Extraction (RPE) Method for 

Detection of Oil Contamination in NonAqueous Drilling Fluids (NAF)

EPA Method 1670

Which can be confirmed by :

Determination of Crude Oil Contamination in 

Non-Aqueous Drilling Fluids by Gas 

Chromatography/Mass Spectrometry (GC/MS) 

EPA Method 1655

§435.13, §435.15

Base fluid retained on 

cuttings

Determination of the Amount of Non-Aqueous 

Drilling Fluid (NAF) Base Fluid from Drill 

Cuttings by a Retort Chamber (Derived from 

API Recommended Practice 13B–2)

EPA Method 1674

§435.13, §435.15
1-3

Table 1-1. Methods by Waste and Pollutant, Subpart A − Offshore Subcategory

Waste source

Regulated Pollutant 

Parameter Analytical/Test Method

Cited in 40 CFR 

Part

Drill cuttings 

associated with nonaqueous drilling 

fluids: Stock 

limitations (C16 –C18

internal olefin)

Mercury 40 CFR Part 136 §435.13, §435.15

Cadmium 40 CFR Part 136 §435.13, §435.15

Polynuclear Aromatic 

Hydrocarbons (PAH)

PAH Content of Oil by HPLC/UV

EPA Method 1654, Revision A

§435.13, §435.15

Base fluid sediment 

toxicity (10-Day Test)

Method for Conducting a Sediment Toxicity 

Test with Leptocheirus Plumulosus and NonAqueous Drilling Fluids or Synthetic-Based 

Drilling Muds (EPA Method 1644)

after using

Procedure for Mixing Base Fluids with 

Sediments

EPA Method 1646

§435.13, §435.15

Biodegradation rate Protocol for the Determination of Degradation 

of Non-Aqueous Base Fluids in a Marine 

Closed Bottle Biodegradation Test System: 

Modified ISO 11734:1995

EPA Method 1647

§435.13, §435.15

Well treatment, 

completion, and 

workover fluids

Oil and grease 40 CFR Part 136 §435.13, §435.15

Free oil Static Sheen Test

EPA Method 1617

§435.12, §435.14

Deck drainage Free oil Visual sheen §435.12, §435.13, 

§435.14, §435.15

Domestic waste Foam Observation §435.13, §435.15

Floating solids Observation §435.14, §435.15

All other domestic 

waste

See 33 CFR part 151 ‡ §435.14, §435.15

Sanitary M10 Total residual chlorine 40 CFR Part 136 §435.12, §435.14, 

§435.15

Sanitary M9IM Floating solids Observation §435.12, §435.14, 

§435.15

§435.12 – Off shore BPT

§435.13 – Offshore BAT

§435.14 – Offshore BCT

§435.15 – Offshore NSPS

† There is no discharge of diesel oil (see §435.13 and §435.15). Diesel oil refers to the grade of distillate fuel oil, as 

specified in ASTM D975-91.

‡ This standard references a U.S. Coast Guard regulation: Vessels Carrying Oil, Noxious Liquid Substances, 

Garbage, Municipal or Commercial Waste, And Ballast Water (33 CFR part 151).

Note: The Offshore Subcategory bans the discharge of the following waste sources and thus there are no related 

analytical or test methods: non-aqueous drilling fluids (NAFs) and produced sand (see §435.12, §435.13, §435.14, 

§435.15).
1-4

Table 1-2. Methods by Waste and Pollutant, Subpart D − Coastal Subcategory

Waste source

Regulated Pollutant 

Parameter Analytical/Test Method

Cited in 40 CFR 

Part

Produced Water Oil & grease 40 CFR Part 136 §435.42, §435.43, 

§435.44, §435.45

Water-based drilling 

fluids, drill cuttings, 

and dewatering 

effluent

SPP Toxicity Drilling Fluids Toxicity Test 

EPA Method 1619

§435.43, §435.45

Free oil Static Sheen Test

EPA Method 1617

§435.42, §435.43, 

§435.44, §435.45

Diesel oil † ASTM specification D975-91 §435.43, §435.45

Mercury 40 CFR Part 136 §435.43, §435.45

Cadmium 40 CFR Part 136 §435.43, §435.45

Well treatment, 

workover, and 

completion fluids

Oil and grease 40 CFR Part 136 §435.43, §435.45

Free oil Static Sheen Test

EPA Method 1617

§435.42, §435.44

Deck drainage Free oil Visual sheen §435.42,§435.43, 

§435.44, §435.45

Domestic waste Foam Observation §435.43, §435.45

Floating solids and 

garbage

Observation §435.42, §435.44, 

§435.45

Sanitary M10 Total residual chlorine 40 CFR Part 136 §435.42, §435.44, 

§435.45

Sanitary M9IM Floating solids Observation §435.42, §435.44, 

§435.45

§435.42 – Coastal BPT

§435.43 – Coastal BAT

§435.44 – Coastal BCT

§435.45 – Coastal NSPS

† There is no discharge of diesel oil (see §435.43 and §435.45). Diesel oil refers to the grade of distillate fuel oil, as 

specified in ASTM D975-91.

Notes:

(1) The Coastal Subcategory bans the discharge of the following waste sources and thus there are no related 

analytical or test methods: non-aqueous drilling fluids (NAFs) and produced sand (see §435.42, §435.43, §435.44, 

§435.45).

(2) The Coastal Subcategory bans the discharge of drill cuttings associated with non-aqueous drilling fluids unless 

there are technical limitations (see §435.43 and §435.45). Appendix 1 of 40 CFR Part 435 Subpart D provides the 

method for permit writers to determine when operators qualify for an exemption from this discharge prohibition. For 

those operators that quality for the exemption from the discharge prohibition EPA requires that these operators meet 

the same stock limitations and discharge limitations for drill cuttings associated with non-aqueous drilling fluids for 

operators in Offshore waters (see 40 CFR §435.13 and §435.15) and no discharge of free oil (determined by the 

Static Sheen Test, required by §435.42, §435.44) in order to discharge drill cuttings associated with non-aqueous 

drilling fluids.
1-5

1.1 Static Sheen Test (EPA Method 1617)

Scope and Application

This method is to be used as a compliance test for the “no discharge of free oil” requirement for 

discharges of drilling fluids, drill cuttings, and well treatment, completion and workover fluids. 

“Free oil” refers to any oil contained in a waste stream that when discharged will cause a film or 

sheen upon or a discoloration of the surface of the receiving water.

Summary of Method

Samples (15-mL) of drilling fluids or well treatment, completion, and workover fluids, and 15-g 

samples (wet weight basis) of drill cuttings or produced sand are introduced into ambient 

seawater in a container having an air-to-liquid interface area of 1,000 cm2(155.5 in2). Samples 

are dispersed within the container and observations made no more than one hour later to 

ascertain if these materials cause sheen, iridescence, gloss, or increased reflectance on the 

surface of the test seawater. The occurrence of any of these visual observations will constitute a 

demonstration that the tested material contains “free oil,” and therefore results in a prohibition of 

its discharge into receiving waters.

1.2 Drilling Fluids Toxicity Test (EPA Method 1619)

Scope and Application

This method is to be used as a compliance test for the suspended particulate phase (SPP) toxicity

requirement for discharges of water-based drilling fluids, associated drill cuttings, and 

dewatering effluent and for drill cuttings associated with non–aqueous drilling fluids. The test 

may be conducted as a full bioassay to estimate the concentration that is lethal to 50% of the test 

organisms (LC50) or as a partial test to determine compliance with 40 CFR Part 435 

requirements. 

Summary of Method

Samples of drilling fluids or drilling mud from active field systems are diluted with natural or 

artificial seawater, mixed, and settled for 1 hour. The suspended particulate phase (SPP) is 

decanted. The decanted solution, defined to be 100% SPP, is further diluted with seawater to 

obtain test concentrations. Mysidopsis bahia (mysid shrimp) are exposed to the test 

concentrations for 96 hours, at which time the living organisms are counted. For the full 

bioassay, a range of concentrations is tested and the LC50 is calculated. For the partial test, a 

negative control, 3% concentration, and positive control are tested. If the number of organisms 

killed in the 3% test concentration meets a number specified in the method, the test material 

passes the partial toxicity test. 

1.3 Procedure for Mixing Base Fluids with Sediments (EPA Method 1646)

This method is used to amend uncontaminated and nontoxic (control) sediments with the base 

fluids that are used to formulate synthetic-based drilling fluids and other non-aqueous drilling 

fluids. Initially, control sediments are press-sieved through a 2,000 micron mesh sieve to remove 

large debris. Then the control sediments are press-sieved through a 500 micron sieve to remove 

indigenous organisms that may prey on the test species or otherwise confound test results. The 

control sediment is homogenized to limit the effects of settling that may have occurred during 

storage. The control sediments are homogenized before density determinations and before the 

addition of base fluid. Because base fluids are strongly hydrophobic and do not readily mix with 
1-6

sediment, care must be taken to ensure base fluids are thoroughly homogenized within the 

sediment. All concentrations are weight-to-weight (mg of base fluid to kg of dry control 

sediment). 

1.4 Protocol for the Determination of Degradation of Non-Aqueous Base Fluids in a 

Marine Closed Bottle Biodegradation Test System: Modified ISO 11734:1995 (EPA 

Method 1647)

Scope and Application

EPA promulgated the use of the marine anaerobic biodegradation analytic method because it 

most closely modeled the ability of a drilling fluid to biodegrade anaerobically in marine 

environments (January 22, 2001; 66 FR 6864). This method determines the anaerobic 

degradation potential of mineral oils, paraffin oils and non-aqueous fluids (NAF) in sediments. 

These substrates are base fluids for formulating offshore drilling fluids. The test evaluates base 

fluid biodegradation rates by monitoring headspace gas production due to microbial degradation 

of the test fluid in natural marine sediment.

Subsequent to this promulgation, EPA incorporated additional quality assurance procedures for 

the marine anaerobic biodegradation analytic method in the NPDES permit for the Western Gulf 

of Mexico (“Final NPDES General Permit for New and Existing Sources and New Dischargers 

in the Offshore Subcategory of the Oil and Gas Extraction Category for the Western Portion of 

the Outer Continental Shelf of the Gulf of Mexico,” GMG290000, Appendix B). The additional 

quality assurance instructions in the GMG290000 more clearly describe the sample preparation 

and compliance determination steps. Specifically, these additional quality assurance procedures 

clarify that users must only use headspace gas to determine compliance with the Part 435 

effluent guidelines. EPA noticed its intention to include these additional quality assurance 

procedures into Part 435 in the “Guidelines Establishing Test Procedures for the Analysis of 

Pollutants Under the Clean Water Act” proposed rulemaking (September 23, 2010; 75 FR 

58023). After responding to public comments EPA included these additional quality assurance 

procedures into Part 435 with the final rule.

Summary of Method

A mixture of marine/estuarine sediment, test substrate (hydrocarbon or controls), and seawater is 

placed into clean 120 ml (150 ml actual volume) Wheaton serum bottles. The test is run using 

four replicate serum bottles containing 2,000 mg carbon/kg dry weight concentration of test 

substrate in sediment. The anaerobic (redox) condition of the bottles is evaluated using resazurin 

dye solution (dye is blue when oxygen is present, reddish in low oxygen conditions and colorless 

if oxygen free). Headspace air is removed with a nitrogen sparge before incubation begins. Gas 

production and composition are measured approximately every two weeks. Gas production is 

measured using a pressure gauge. Barometric pressure is measured at the time of testing to make 

necessary volume adjustments. The test period is 275 days. The results of EPA Method 1647 for 

the test fluid are compared against the reference fluid to determine whether the test fluid is 

eligible for discharge with drill cuttings.
1-7

1.5 Determination of Crude Oil Contamination in Non-Aqueous Drilling Fluids by Gas 

Chromatography/Mass Spectrometry (GC/MS) (EPA Method 1655)

Scope and Application

This method determines crude (formation) oil contamination, or other petroleum oil 

contamination, in non-aqueous drilling fluids (NAFs) by comparing the gas 

chromatography/mass spectrometry (GC/MS) fingerprint scan and extracted ion scans of the test 

sample to that of an uncontaminated sample. It can be used for monitoring oil contamination of 

NAFs or monitoring oil contamination of the base fluid used in the NAF formulations.

Summary of Method

Analysis of NAF for crude oil contamination is a step-wise process. The analyst first performs a 

qualitative assessment of the presence or absence of crude oil in the sample. If crude oil is 

detected during this qualitative assessment, the analyst must perform a quantitative analysis of 

the crude oil concentration. A sample of NAF is centrifuged to obtain a solids free supernate.

The test sample is prepared by removing an aliquot of the solids free supernate, spiking it with 

internal standard, and analyzing it using GC/MS techniques. The components are separated by 

the gas chromatograph and detected by the mass spectrometer.

Qualitative identification of crude oil contamination is performed by comparing the Total Ion 

Chromatograph (TIC) scans and Extracted Ion Profile (EIP) scans of test sample to that of 

uncontaminated base fluids, and examining the profiles for chromatographic signatures 

diagnostic of oil contamination. The presence or absence of crude oil contamination observed in 

the full scan profiles and selected extracted ion profiles determines further sample quantitation 

and reporting requirements. If crude oil is detected in the qualitative analysis, quantitative 

analysis must be performed by calibrating the GC/MS using a designated NAF spiked with 

known concentrations of a designated oil.

1.6 Reverse Phase Extraction (RPE) Method for Detection of Oil Contamination in NonAqueous Drilling Fluids (NAF) (EPA Method 1670)

Scope and Application

This method is used for determination of crude or formation oil, or other petroleum oil 

contamination, in non-aqueous drilling fluids (NAFs). It is intended as a positive/negative test to 

determine a presence of crude oil in NAF prior to discharging drill cuttings from offshore 

production platforms. It is for use in the Environmental Protection Agency's (EPA's) survey and 

monitoring programs under the Clean Water Act, including monitoring of compliance with the 

Gulf of Mexico NPDES General Permit for monitoring of oil contamination in drilling fluids.

The method has been designed to show positive contamination for 5% of representative crude 

oils at a concentration of 0.1% in drilling fluid (vol/vol), 50% of representative crude oils at a 

concentration of 0.5%, and 95% of representative crude oils at a concentration of 1%.

Summary of Method

An aliquot of drilling fluid is extracted using isopropyl alcohol. The mixture is allowed to settle 

and then filtered to separate out residual solids. An aliquot of the filtered extract is charged onto 

a reverse phase extraction (RPE) cartridge. The cartridge is eluted with isopropyl alcohol. Crude 

oil contaminates are retained on the cartridge and their presence (or absence) is detected based 

on observed fluorescence using a black light.
1-8

1.7 Determination of the Amount Of Non-Aqueous Drilling Fluid (NAF) Base Fluid from 

Drill Cuttings by a Retort Chamber (Derived From API Recommended Practice 13B–

2) (EPA Method 1674)

Scope and Application

This procedure is specifically intended to measure the amount of non-aqueous drilling fluid 

(NAF) base fluid from cuttings generated during a drilling operation. This procedure is a retort 

test which measures all oily material (NAF base fluid) and water released from a cuttings sample 

when heated in a calibrated and properly operating “Retort” instrument.

Summary of Method

A known mass of cuttings is heated in the retort chamber to vaporize the liquids associated with 

the sample. The NAF base fluid and water vapors are then condensed, collected, and measured in 

a precision graduated receiver.

1.8 PAH Content of Oil by HPLC/UV (EPA Method 1654, Revision A)

Scope and Application

This method is designed to determine the polynuclear aromatic hydrocarbon (PAH) content of 

oil by high-performance liquid chromatography (HPLC) with an ultra-violet absorption (UV) 

detector. The PAH content is measured and reported as phenanthrene. For oil in drilling muds, 

this method is designed to be used in conjunction with the extraction procedure in EPA Method 

1662.

Summary of Method

An oil sample is diluted in acetonitrile and a 20-μL aliquot is injected into the HPLC. The

PAHs are partially separated by HPLC and detected with the UV detector. Identification of PAH 

(qualitative analysis) is performed by comparing the response of the UV detector to the response 

during the retention-time range characteristic of the PAH in diesel oil. PAH is present when a 

response occurs during this retention-time range. Quantitative analysis is performed by 

calibrating the HPLC with phenanthrene using an external standard technique, and using the 

calibration factor to determine the concentration of PAH in the sample.

1.9 Sediment Toxicity Test for NAF and SBM (EPA Method 1644)

Scope and Application

This test method describes the procedures for obtaining data regarding the effects of nonaqueous drilling fluids (NAFs) or synthetic based drilling muds (SBMs) on the marine 

amphipod, Leptocheirus plumulosus. EPA regulates the sediment toxicity in NAFs and SBMs 

that are discharged from oil and gas extraction facilities in coastal and offshore waters as an 

indicator for toxic pollutants (see 40 CFR 435.13, 435.15, 435.43, and 435.45). EPA established 

the use of this test and related limits to encourage the use of less toxic drilling fluids and 

additives. The sediment toxicity of the NAF-cuttings at the point of discharge is measured by

this modified sediment toxicity test using a natural sediment and Leptocheirus plumulosus as the 

test organism.

EPA promulgated the use of this method with revisions to the Oil and Gas Extraction effluent 

guidelines (January 22, 2001; 66 FR 6849). This initial method was consistent with ASTM 

Standard Guide E 1367-92 (ASTM 1997). Subsequent to this rulemaking EPA updated this 
1-9

method in the NPDES permit for the Western Gulf of Mexico (GMG290000, Appendix A) to be 

consistent with ASTM E 1367-99. Section 10 of this document re-prints the sediment toxicity

test from the NPDES permit for the Western Gulf of Mexico (GMG290000, Appendix A).

Summary of Method

Samples of NAFs and SBMs are exposed to cultured Leptocheirus plumulosus test organisms. 

This method relies on standardized bioassay benthic toxicity procedures from ASTM E 1367 and 

EPA/600/R-94/0258 along with modified sediment mixing procedures that address the unique 

properties of SBM (EPA Method 1646). Of the several species typically used for these tests, 

Leptocheirus plumulosus was selected because it demonstrated the range of sensitivity to the 

base fluids that were under evaluation and provided both repeatability and discriminatory power. 

L. plumulosus live in fine-grained sediments and warm waters similar to the conditions found in 

the anticipated receiving environment of the Gulf of Mexico. L. plumulosus is also easy to 

culture in the lab and therefore available year-round for testing. The only SBM drill cuttings that 

can be discharge are cuttings coated with NAFs and SBM that are as toxic or less toxic, but not 

more toxic, than the reference NAF and SBM (C16-C18 internal olefin or C12-C14 or C8 ester) as 

measured by LC50. This method tests the whole SBM (96-hour exposure test, discharge 

limitation) and the NAF (10-day exposure test, stock limitations). This method is referenced in 

40 CFR 435.13, 435.15, 435.43, and 435.45. 

1.10 Previous Publication of Oil and Gas Extraction Point Source Category Analytic 

Methods

EPA previously published the analytical methods for the Oil and Gas Extraction Point Source 

Category in the Code of Federal Regulations (CFR). These methods were published as 

attachments to Subpart A (Offshore Subcategory). Table 1-3 provides a listing of these methods 

as they appear in the CFR and their current EPA method number. 

Subsequent to these rulemakings EPA proposed several changes to Part 435, Oil and Gas 

Extraction Point Source Category on 23 September 2010 (75 FR 58023). EPA solicited comment 

on moving, and in two cases revising, the methods from 40 CFR Part 435, Subpart A (Offshore 

Subcategory) to this EPA document, which was included in the rulemaking record for the 

proposed rule.1 EPA proposed to organize the analytical methods for the Offshore Subcategory

into this document in order to allow for easier access to the methods for this category. EPA also 

proposed to incorporate additional quality assurance procedures in the marine anaerobic 

biodegradation analytic method (Appendix 4 to Subpart A of Part 435) and to correct some 

erroneous references and omissions in the method for identification of crude oil contamination

(Appendix 5 to Subpart A of Part 435). EPA noticed its intent to include these revisions to these 

two methods in the previous version of this document. 

EPA received three sets of comments on the above-proposed actions. EPA reviewed and 

responded to these comments (see EPA-HQ-OW-2010-0192-0101, EPA-HQ-OW-2010-0192-

0155, EPA-HQ-OW-2010-0192-0127, and the Response to Comments Document, Docket No. 

EPA-HQ-OW-2010-0192). In the final rulemaking (2011) EPA retained the full publication of 

the Oil and Gas Extraction analytical methods (Appendix 1 through 7 to Subpart A) from 40 

 

1

See “Guidelines Establishing Test Procedures for the Analysis of Pollutants Under the Clean Water Act; Analysis 

and Sampling Procedures,” RIN 2040-AF09, Docket No. EPA-HQ-OW-2010-0192-0034. Available at: 

http://www.epa.gov/regulations.
1-10

CFR 435, Subpart A, and incorporated these methods by reference to this document.2Like any

other changes to an EPA approved method, any changes to the methods in this document will 

require a rulemaking.

Table 1-3. EPA Method Numbers for Oil and Gas Extraction Point Source Category

Analytical Methods and Prior CFR References in 40 CFR 435

Analytical/Test Method

EPA Method 

Number

Date First 

Promulgated CFR References

Static Sheen Test 1617 1993 Subpart A, Appendix 1

Drilling Fluids Toxicity Test 1619 1993 Subpart A, Appendix 2

Procedure for Mixing Base Fluids With Sediments 1646 2001 Subpart A, Appendix 3

Protocol for the Determination of Degradation of 

Non-Aqueous Base Fluids in a Marine Closed Bottle 

Biodegradation Test System: Modified ISO 

11734:1995

1647 2001 Subpart A, Appendix 4

Determination of Crude Oil Contamination in NonAqueous Drilling Fluids by Gas 

Chromatography/Mass Spectrometry (GC/MS)

1655 2001 Subpart A, Appendix 5

Reverse Phase Extraction (RPE) Method for Detection 

of Oil Contamination in Non-Aqueous Drilling Fluids 

(NAF)

1670 2001 Subpart A, Appendix 6

Determination of the Amount of Non-Aqueous 

Drilling Fluid (NAF) Base Fluid from Drill Cuttings 

by a Retort Chamber (Derived from API 

Recommended Practice 13B–2)

1674 2001 Subpart A, Appendix 7

Method for Conducting a Sediment Toxicity Test with 

Leptocheirus Plumulosus and Non-Aqueous Drilling 

Fluids or Synthetic-Based Drilling Muds

1644 2001 N/A †

† Note: EPA Method 1644 (sediment toxicity test) was first promulgated for use in 2001 but was incorporated by 

reference. Subsequent to this rulemaking EPA updated this method in the NPDES permit for the Western Gulf of 

Mexico (GMG290000, Appendix A) to be consistent with ASTM E 1367-99 (ASTM 2000). Section 10 of this 

document re-prints the sediment toxicity test from the NPDES permit for the Western Gulf of Mexico 

(GMG290000, Appendix A).

 

2

 Federal regulations at 1 CFR 51.7 (c)(1) prohibit the incorporation by reference of material published previously 

in the Federal Register. These regulations prevent EPA from deleting these methods from 40 CFR 435.
2-11

2. STATIC SHEEN TEST (EPA METHOD 1617)

2.1 Scope and Application

This method is to be used as a compliance test for the “no discharge of free oil” requirement for 

discharges of drilling fluids, drill cuttings, produced sand, and well treatment, completion and 

workover fluids. “Free oil” refers to any oil contained in a waste stream that when discharged 

will cause a film or sheen upon or a discoloration of the surface of the receiving water.

2.2 Summary of Method

15-mL samples of drilling fluids or well treatment, completion, and workover fluids, and 15-g 

samples (wet weight basis) of drill cuttings or produced sand are introduced into ambient 

seawater in a container having an air-to-liquid interface area of 1,000 cm2(155.5 in2). Samples 

are dispersed within the container and observations made no more than one hour later to 

ascertain if these materials cause a sheen, iridescence, gloss, or increased reflectance on the 

surface of the test seawater. The occurrence of any of these visual observations will constitute a 

demonstration that the tested material contains “free oil,” and therefore results in a prohibition of 

its discharge into receiving waters.

2.3 Interferences

Residual “free oil” adhering to sampling containers, the magnetic stirring bar used to mix the 

sample, and the stainless steel spatula used to mix the sample will be the principal sources of 

contamination problems. These problems should only occur if improperly washed and cleaned 

equipment are used for the test. The use of disposable equipment minimizes the potential for 

similar contamination from pipettes and the test container.

2.4 Apparatus, Materials, and Reagents

2.4.1 Apparatus

2.4.1.1 Sampling Containers: 1-liter polyethylene beakers and 1-liter glass 

beakers.

2.4.1.2 Graduated cylinder: 100-mL graduated cylinder required only for 

operations where predilution of mud discharges is required.

2.4.1.3 Plastic disposable weighing boats.

2.4.1.4 Triple-beam scale.

2.4.1.5 Disposable pipettes: 25-mL disposable pipettes.

2.4.1.6 Magnetic stirrer and stirring bar.

2.4.1.7 Stainless steel spatula.
2-12

2.4.1.8 Test container: Open plastic container whose internal cross-section 

parallel to its opening has an area of 1,000 cm2±50 cm2 (155.5 ±7.75 

in2), and a depth of at least 13 cm (5 inches) and no more than 30 cm (11.8 

inches).

2.4.2 Materials and Reagents.

2.4.2.1 Plastic liners for the test container: Oil-free, heavy-duty plastic trash can 

liners that do not inhibit the spreading of an oil film. Liners must be of 

sufficient size to completely cover the interior surface of the test container. 

Permittees must determine an appropriate local source of liners that do not 

inhibit the spreading of 0.05 mL of diesel fuel added to the lined test 

container under the test conditions and protocol described below.

2.4.2.2 Ambient receiving water.

2.5 Calibration

None currently specified.

2.6 Quality Control Procedures

None currently specified.

2.7 Sample Collection and Handling

2.7.1 Sampling containers must be thoroughly washed with detergent, rinsed a minimum of 

three times with fresh water, and allowed to air dry before samples are collected.

2.7.2 Samples of drilling fluid to be tested shall be taken at the shale shaker after cuttings 

have been removed. The sample volume should range between 200 mL and 500 mL.

2.7.3 Samples of drill cuttings will be taken from the shale shaker screens with a clean 

spatula or similar instrument and placed in a glass beaker. Cuttings samples shall be 

collected prior to the addition of any washdown water and should range between 

200 g and 500 g.

2.7.4 Samples of produced sand must be obtained from the solids control equipment from 

which the discharge occurs on any given day and shall be collected prior to the 

addition of any washdown water; samples should range between 200 g and 500 g.

2.7.5 Samples of well treatment, completion, and workover fluids must be obtained from 

the holding facility prior to discharge; the sample volume should range between 

200 mL and 500 mL.

2.7.6 Samples must be tested no later than 1 hour after collection.

2.7.7 Drilling fluid samples must be mixed in their sampling containers for 5 minutes prior 

to the test using a magnetic bar stirrer. If predilution is imposed as a permit condition, 
2-13

the sample must be mixed at the same ratio with the same prediluting water as the 

discharged muds and stirred for 5 minutes.

2.7.8 Drill cuttings must be stirred and well mixed by hand in their sampling containers 

prior to testing, using a stainless steel spatula.

2.8 Procedure

2.8.1 Ambient receiving water must be used as the “receiving water” in the test. The 

temperature of the test water shall be as close as practicable to the ambient conditions 

in the receiving water, not the room temperature of the observation facility. The test 

container must have an air-to-liquid interface area of 1,000 ±50 cm2. The surface of 

the water should be no more than 1.27 cm (0.5 inch) below the top of the test 

container.

2.8.2 Plastic liners shall be used, one per test container, and discarded afterwards. Some 

liners may inhibit spreading of added oil; operators shall determine an appropriate 

local source of liners that do not inhibit the spreading of the oil film.

2.8.3 A 15-mL sample of drilling fluid or well treatment, completion, and workover fluids 

must be introduced by pipette into the test container 1 cm below the water surface. 

Pipettes must be filled and discharged with test material prior to the transfer of test 

material and its introduction into test containers. The test water/test material mixture 

must be stirred using the pipette to distribute the test material homogeneously

throughout the test water. The pipette must be used only once for a test and then 

discarded.

2.8.4 Drill cuttings or produced sand should be weighed on plastic weighing boats; 15-g 

samples must be transferred by scraping test material into the test water with a 

stainless steel spatula. Drill cuttings shall not be prediluted prior to testing. Also, 

drilling fluids and cuttings will be tested separately. The weighing boat must be 

immersed in the test water and scraped with the spatula to transfer any residual 

material to the test container. The drill cuttings or produced sand must be stirred with 

the spatula to an even distribution of solids on the bottom of the test container.

2.8.5 Observations must be made no later than 1 hour after the test material is transferred to 

the test container. Viewing points above the test container should be made from at 

least three sides of the test container, at viewing angles of approximately 60° and 30° 

from the horizontal. Illumination of the test container must be representative of 

adequate lighting for a working environment to conduct routine laboratory

procedures. It is recommended that the water surface of the test container be observed 

under a fluorescent light source such as a dissecting microscope light. The light 

source shall be positioned above and directed over the entire surface of the pan.

2.8.6 Detection of a “silvery” or “metallic” sheen or gloss, increased reflectivity, visual 

color, iridescence, or an oil slick on the water surface of the test container surface 

shall constitute a demonstration of “free oil.” These visual observations include 

patches, streaks, or sheets of such altered surface characteristics. If the free oil content 

of the sample approaches or exceeds 10%, the water surface of the test container may
2-14

lack color, a sheen, or iridescence, due to the increased thickness of the film; thus, the 

observation for an oil slick is required. The surface of the test container shall not be 

disturbed in any manner that reduces the size of any sheen or slick that may be 

present.

If an oil sheen or slick occurs on less than one-half of the surface area after the 

sample is introduced to the test container, observations will continue for up to 1 hour. 

If the sheen or slick increases in size and covers greater than one-half of the surface 

area of the test container during the observation period, the discharge of the material 

shall cease. If the sheen or slick does not increase in size to cover greater than onehalf of the test container surface area after one hour of observation, discharge may

continue and additional sampling is not required.

If a sheen or slick occurs on greater than one-half of the surface area of the test 

container after the test material is introduced, discharge of the tested material shall 

cease. The permittee may retest the material causing the sheen or slick. If subsequent 

tests do not result in a sheen or slick covering greater than one-half of the surface area 

of the test container, discharge may continue.
3-1

3. DRILLING FLUIDS TOXICITY TEST (EPA METHOD 1619)

3.1 Scope and Application

This method is to be used as a compliance test for the suspended particulate phase (SPP) toxicity

requirement for discharges of water-based drilling fluids, associated drill cuttings, and 

dewatering effluent and for drill cuttings associated with non –aqueous drilling fluids. The test 

may be conducted as a full bioassay to estimate the concentration that is lethal to 50% of the test 

organisms (LC50) or as a partial test to determine compliance with 40 CFR Part 435 

requirements. 

3.2 Summary of Method

Samples of drilling fluids or drilling mud from active field systems are diluted with natural or 

artificial seawater, mixed, and settled for 1 hour. The suspended particulate phase (SPP) is 

decanted. The decanted solution, defined to be 100% SPP, is further diluted with seawater to 

obtain test concentrations. Mysidopsis bahia (mysid shrimp) are exposed to the test 

concentrations for 96 hours, at which time the living organisms are counted. For the full 

bioassay, a range of concentrations is tested and the LC50 is calculated. For the partial test, a 

negative control, 3% concentration, and positive control are tested. If the number of organisms 

killed in the 3% test concentration meets a number specified in the method, the test material 

passes the partial toxicity test. 

3.3 Sample Collection

The collection and preservation methods for drilling fluids (muds) and water samples presented 

here are designed to minimize sample contamination and alteration of the physical or chemical 

properties of the samples due to freezing, air oxidation, or drying.

3.3.1 Apparatus

3.3.1.1 The following items are required for water and drilling mud sampling and 

storage:

a. Acid-rinsed linear-polyethylene bottles or other appropriate 

noncontaminating drilling mud sampler.

b. Acid-rinsed linear-polyethylene bottles or other appropriate 

noncontaminating water sampler.

c. Acid-rinsed linear-polyethylene bottles or other appropriate 

noncontaminated vessels for water and mud samples.

d. Ice chests for preservation and shipping of mud and water samples.

3.3.2 Water Sampling

3.3.2.1 Collection of water samples shall be made with appropriate acid-rinsed 

linear-polyethylene bottles or other appropriate non-contaminating water 

sampling devices. Special care shall be taken to avoid the introduction of 
3-2

contaminants from the sampling devices and containers. Prior to use, the 

sampling devices and containers should be thoroughly cleaned with a 

detergent solution, rinsed with tap water, soaked in 10% hydrochloric acid 

(HCl) for 4 hours, and then thoroughly rinsed with glass-distilled water.

3.3.3 Drilling Mud Sampling

3.3.3.1 Drilling mud formulations to be tested shall be collected from active field 

systems. Obtain a well-mixed sample from beneath the shale shaker after 

the mud has passed through the screens. Samples shall be stored in 

polyethylene containers or in other appropriate uncontaminated vessels. 

Prior to sealing the sample containers on the platform, flush as much air 

out of the container by filling it with drilling fluid sample, leaving a one 

inch space at the top.

3.3.3.2 Mud samples shall be immediately shipped to the testing facility on blue 

or wet ice (do not use dry ice) and continuously maintained at 0–4°C until 

the time of testing.

3.3.3.3 Bulk mud samples shall be thoroughly mixed in the laboratory using a 

1,000 rpm high shear mixer and then subdivided into individual, small 

wide-mouthed (e.g., one or two liter) non-contaminating containers for 

storage.

3.3.3.4 The drilling muds stored in the laboratory shall have any excess air 

removed by flushing the storage containers with nitrogen under pressure 

anytime the containers are opened. Moreover, the sample in any container 

opened for testing must be thoroughly stirred using a 1,000 rpm high shear 

mixer prior to use.

3.3.3.5 Most drilling mud samples may be stored for periods of time longer than 2 

weeks prior to toxicity testing provided that proper containers are used and 

proper condition are maintained.

3.4 Suspended Particulate Phase Sample Preparation

Mud samples that have been stored under specified conditions in this protocol shall be prepared 

for tests within three months after collection. The SPP shall be prepared as detailed below.

3.4.1 Apparatus

3.4.1.1 The following items are required:

a. Magnetic stir plates and bars.

b. Several graduated cylinders, ranging in volume from 10 mL to 1 L

c. Large (15 cm) powder funnels.

d. Several 2-liter graduated cylinders.
3-3

e. Several 2-liter large mouth graduated Erlenmeyer flasks.

3.4.1.2 Prior to use, all glassware shall be thoroughly cleaned. Wash all glassware 

with detergent, rinse five times with tap water, rinse once with acetone, 

rinse several times with distilled or deionized water, place in a clean 10% 

(or stronger) HCl acid bath for a minimum of 4 hours, rinse five times 

with tap water, and then rinse five times with distilled or deionized water. 

For test samples containing mineral oil or diesel oil, glassware should be 

washed with petroleum ether to assure removal of all residual oil.

Note: If the glassware with nytex cups soaks in the acid solution longer 

than 24 hours, then an equally long deionized water soak should be 

performed.

3.4.2 Test Seawater Sample Preparation

3.4.2.1 Diluent seawater and exposure seawater samples are prepared by filtration 

through a 1.0 micrometer filter prior to analysis.

3.4.2.2 Artificial seawater may be used as long as the seawater has been prepared 

by standard methods or ASTM methods, has been properly “seasoned,” 

filtered, and has been diluted with distilled water to the same specified 

20 ±2 ppt salinity and 20 ±2°C temperature as the “natural” seawater.

3.4.3 Sample Preparation

3.4.3.1 The pH of the mud shall be tested prior to its use. If the pH is less than 9, 

if black spots have appeared on the walls of the sample container, or if the 

mud sample has a foul odor, that sample shall be discarded. Subsample a 

manageable aliquot of mud from the well-mixed original sample. Mix the 

mud and filtered test seawater in a volumetric mud-to-water ratio of 1 to 9. 

This is best done by the method of volumetric displacement in a 2–L, large 

mouth, graduated Erlenmeyer flask. Place 1,000 mL of seawater into the 

graduated Erlenmeyer flask. The mud subsample is then carefully added 

via a powder funnel to obtain a total volume of 1200 mL. (A 200 mL 

volume of the mud will now be in the flask).

The 2–L, large mouth, graduated Erlenmeyer flask is then filled to the 

2,000 mL mark with 800 mL of seawater, which produces a slurry with a 

final ratio of one volume drilling mud to nine volumes water. If the 

volume of SPP required for testing or analysis exceeds 1,500 to 1,600 mL, 

the initial volumes should be proportionately increased. Alternatively, 

several 2–L drill mud/water slurries may be prepared as outlined above 

and combined to provide sufficient SPP.

3.4.3.2 Mix this mud/water slurry with magnetic stirrers for 5 minutes. Measure 

the pH and, if necessary, adjust (decrease) the pH of the slurry to within 

0.2 units of the seawater by adding 6N HCl while stirring the slurry. Then, 

allow the slurry to settle for 1 hour. Record the amount of HCl added.
3-4

3.4.3.3 At the end of the settling period, carefully decant (do not siphon) the 

Suspended Particulate Phase (SPP) into an appropriate container. 

Decanting the SPP is one continuous action. In some cases no clear 

interface will be present; that is, there will be no solid phase that has 

settled to the bottom. For those samples the entire SPP solution should be 

used when preparing test concentrations. However, in those cases when no 

clear interface is present, the sample must be remixed for five minutes. 

This insures the homogeneity of the mixture prior to the preparation of the 

test concentrations. In other cases, there will be samples with two or more 

phases, including a solid phase. For those samples, carefully and 

continuously decant the supernatant until the solid phase on the bottom of 

the flask is reached. The decanted solution is defined to be 100% SPP. 

Any other concentration of SPP refers to a percentage of SPP that is 

obtained by volumetrically mixing 100% SPP with seawater.

3.4.3.4 SPP samples to be used in toxicity tests shall be mixed for 5 minutes and 

must not be preserved or stored.

3.4.3.5 Measure the filterable and unfilterable residue of each SPP prepared for 

testing. Measure the dissolved oxygen (DO) and pH of the SPP. If the DO 

is less than 4.9 ppm, aerate the SPP to at least 4.9 ppm which is 65% of 

saturation. Maximum allowable aeration time is 5 minutes using a generic 

commercial air pump and air stone. Neutralize the pH of the SPP to a pH 

7.8 ±0.1 using a dilute HCl solution. If too much acid is added to lower the 

pH saturated NaOH may be used to raise the pH to 7.8 ±0.1 units. Record 

the amount of acid or NaOH needed to lower/raise to the appropriate pH. 

Three repeated DO and pH measurements are needed to insure 

homogeneity and stability of the SPP. Preparation of test concentrations 

may begin after this step is complete.

3.4.3.6 Add the appropriate volume of 100% SPP to the appropriate volume of 

seawater to obtain the desired SPP concentration. The control is seawater 

only. Mix all concentrations and the control for 5 minutes by using 

magnetic stirrers. Then, the animals shall be randomly selected and placed 

in the dishes in order to begin the 96-hour toxicity test.

3.5 Guidance for Performing Suspended Particulate Phase Toxicity Tests Using 

Mysidopsis bahia

3.5.1 Apparatus

3.5.1.1 Each definitive test consists of 18 test containers: 3 replicates of a control 

and 5 SPP dilutions. Test containers should be Pyrex or equivalent glass. 

For definitive tests, 5 SPP dilutions with 3 replicates of at least 500 ml 

each are required. Twenty mysids per replicate, 360 per definitive test are 

required.

3.5.1.2 A range-finding test consists of at least 6 test containers: 1 replicate of a 

control and 5 SPP dilutions. The duration of range-finding tests may be 
3-5

less than 96 hours. Ten or more mysids per replicate, at least 60 per rangefinding test, are suggested. Daily water quality measurements and 

observations are optional with range-finding tests.

3.5.2 Sample Collection Preservation

3.5.2.1 Drilling muds and water samples are collected and stored as described in 

Section 3.3.3.

3.5.3 Species Selection

3.5.3.1 The Suspended Particulate Phase (SPP) tests on drilling muds shall utilize 

the test species Mysidopsis bahia. Test animals shall be 3 to 6 days old on 

the first day of exposure. Whatever the source of the animals, collection 

and handling should be as gentle as possible. Transportation to the 

laboratory should be in well-aerated water from the animal culture site at 

the temperature and salinity from which they were cultured. Methods for 

handling, acclimating, and sizing bioassay organisms given by Borthwick 

[1] and Nimmo [2] shall be followed in matters for which no guidance is 

given here.

3.5.4 Experimental Conditions

3.5.4.1 Suspended particulate phase (SPP) tests should be conducted at a salinity

of 20 ±2 ppt. Experimental temperature should be 20 ±2°C. Dissolved 

oxygen in the SPP shall be raised to or maintained above 65% of 

saturation prior to preparation of the test concentrations. Under these 

conditions of temperature and salinity, 65% saturation is a DO of 5.3 ppm. 

Beginning at Day 0-before the animals are placed in the test containers 

DO, temperature, salinity, and pH shall be measured every 24 hours. DO 

should be reported in milligrams per liter.

3.5.4.2 Aeration of test media is required during the entire test with a rate 

estimated to be 50–140 cubic centimeters/minute. This air flow to each 

test dish may be achieved through polyethylene tubing (0.045-inch inner 

diameter and 0.062-inch outer diameter) by a small generic aquarium 

pump. The delivery method, surface area of the aeration stone, and flow 

characteristics shall be documented. All treatments, including control, 

shall be the same.

3.5.4.3 Light intensity shall be 1,200 microwatts/cm2using cool white fluorescent 

bulbs with a 14-hr light and 10-hr dark cycle. This light/dark cycle shall 

also be maintained during the acclimation period and the test.

3.5.5 Experimental Procedure

3.5.5.1 Wash all glassware with detergent, rinse five times with tap water, rinse 

once with acetone, rinse several times with distilled or deionized water, 
3-6

place in a clean 10% HCl acid bath for a minimum of 4 hours, rinse five 

times with tap water, and then rinse five times with distilled water.

3.5.5.2 Establish the definitive test concentration based on results of a range 

finding test or based on prior experience and knowledge of the mud 

system. A minimum of five test concentrations plus a negative and 

positive (reference toxicant) control is required for the definitive test. To 

estimate the LC50, two concentrations shall be chosen that give (other than 

zero and 100%) mortality above and below 50%.

3.5.5.3 Twenty organisms are exposed in each test dish. Nytex® cups shall be 

inserted into every test dish prior to adding the animals. These “nylon 

mesh screen” nytex holding cups are fabricated by gluing a collar of 363-

micrometer mesh nylon screen to a 15-centimeter wide Petri dish with 

silicone sealant. The nylon screen collar is approximately 5 centimeters 

high. The animals are then placed into the test concentration within the 

confines of the Nytex cups.

3.5.5.4 Individual organisms shall be randomly assigned to treatment. A 

randomization procedure is presented in Section 3.7. Make every attempt 

to expose animals of approximately equal size. The technique described 

by Borthwick [1], or other suitable substitutes, should be used for 

transferring specimens. Throughout the test period, mysids shall be fed 

daily with approximately 50 Artemia (brine shrimp) nauplii per mysid. 

This will reduce stress and decrease cannibalism.

3.5.5.5 Cover the dishes, aerate, and incubate the test containers in an appropriate 

test chamber. Positioning of the test containers holding various 

concentrations of test solution should be randomized if incubator 

arrangement indicates potential position difference. The test medium is not 

replaced during the 96-hour test.

3.5.5.6 Observations may be attempted at 4, 6 and 8 hours; they must be 

attempted at 0, 24, 48, and 72 hours and must be made at 96 hours. 

Attempts at observations refers to placing a test dish on a light table and 

visually counting the animals. Do not lift the “nylon mesh screen” cup out 

of the test dish to make the observation. No unnecessary handling of the 

animals should occur during the 96 hour test period. DO and pH 

measurements must also be made at 0, 24, 48, 72, and 96 hours. Take and 

replace the test medium necessary for the DO and pH measurements 

outside of the nytex cups to minimize stresses on the animals.

3.5.5.7 At the end of 96 hours, all live animals must be counted. Death is the end 

point, so the number of living organisms is recorded. Death is determined 

by lack of spontaneous movement. All crustaceans molt at regular 

intervals, shedding a complete exoskeleton. Care should be taken not to 

count an exoskeleton. Dead animals might decompose or be eaten between 

observations. Therefore, always count living, not dead animals. If daily

observations are made, remove dead organisms and molted exoskeletons 
3-7

with a pipette or forceps. Care must be taken not to disturb living 

organisms and to minimize the amount of liquid withdrawn.

3.6 Methods for Positive Control Tests (Reference Toxicant)

3.6.1.1 Sodium lauryl sulfate (dodecyl sodium sulfate) is used as a reference 

toxicant for the positive control. The chemical used should be 

approximately 95% pure. The source, lot number, and percent purity shall 

be reported.

3.6.1.2 Test methods are those used for the drilling fluid tests, except that the test 

material was prepared by weighing one gram sodium lauryl sulfate on an 

analytical balance, adding the chemical to a 100-milliliter volumetric 

flask, and bringing the flask to volume with deionized water. After mixing 

this stock solution, the test mixtures are prepared by adding 0.1 milliliter 

of the stock solution for each part per million desired to one liter of 

seawater.

3.6.1.3 The mixtures are stirred briefly, water quality is measured, animals are 

added to holding cups, and the test begins. Incubation and monitoring 

procedures are the same as those for the drilling fluids.

3.7 Randomization Procedure

3.7.1 Purpose and Procedure

3.7.1.1 The purpose of this procedure is to assure that mysids are impartially 

selected and randomly assigned to six test treatments (five drilling fluid or 

reference toxicant concentrations and a control) and impartially counted at 

the end of the 96-hour test. Thus, each test setup, as specified in the 

randomization procedure, consists of 3 replicates of 20 animals for each of 

the six treatments, i.e., 360 animals per test. Figure 3-1 is a flow diagram 

that depicts the procedure schematically and should be reviewed to 

understand the over-all operation. The following tasks shall be performed 

in the order listed.

3.7.1.2 Mysids are cultured in the laboratory in appropriate units. If mysids are 

purchased, go to Task 3 (Section 3.7.1.3).

3.7.1.3 Remove mysids from culture tanks (6, 5, 4, and 3 days before the test will 

begin, i.e., Tuesday, Wednesday, Thursday, and Friday if the test will 

begin on Monday) and place them in suitably large maintenance 

containers so that they can swim about freely and be fed.

Note: Not every detail (the definition of suitably large containers, for 

example) is provided here. Training and experience in aquatic animal 

culture and testing will be required to successfully complete these tests.
3-8

1 Culture Units

2 Maintenance 

Containers

3 Test 

Population 

Containers

4 Separation/

Enumeration 

Containers

5 Counting Dish 

(repeat tasks 

5-7 for A1 and 

A2 containers)

6 Distribution 

Containers

7 Test 

Containers

A1 A2 B1 B2

1A 1B 1C 1D

1 2 3 4 5 6 36

6D

If Mysids are 

purchased

Mysids are 

collected 3 to 

6 days prior 

to testing

Task

. . .

. . .

Figure 3-1. Mysid Randomization Procedure

3.7.1.4 Remove mysids from maintenance containers and place all animals in a 

single container. The intent is to have homogeneous test population of

mysids of a known age (3–6 days old).

3.7.1.5 For each toxicity test, assign two suitable containers (500-milliliter (mL) 

beakers are recommended) for mysid separation/enumeration. Label each 

container (A1, A2, B1, B2, and C1, C2, for example, if two drilling fluid 

tests and a reference toxicant test are to be set up on one day). The purpose 

of this task is to allow the investigator to obtain a close estimate of the 

number of animals available for testing and to prevent unnecessary

crowding of the mysids while they are being counted and assigned to test 

containers. Transfer the mysids from the large test population container to 

the labeled separation and enumeration containers but do not place more 

than 200 mysids in a 500-mL beaker. Be impartial in transferring the 
3-9

mysids; place approximately equal numbers of animals (10–15 mysids is 

convenient) in each container in a cyclic manner rather than placing the 

maximum number each container at one time.

Note: It is important that the animals not be unduly stressed during this 

selection and assignment procedure. Therefore, it will probably be 

necessary to place all animals (except the batch immediately being 

assigned to test containers) in mesh cups with flowing seawater or in large 

volume containers with aeration. The idea is to provide the animals with 

near optimal conditions to avoid additional stress.

3.7.1.6 Place the mysids from the two labeled enumeration containers assigned to 

a specific test into one or more suitable containers to be used as counting 

dishes (2-liter Carolina dishes are suggested). Because of the time required 

to separate, count, and assign mysids, two or more people may be 

involved in completing this task. If this is done, two or more counting 

dishes may be used, but the investigator must make sure that 

approximately equal numbers of mysids from each labeled container are 

placed in each counting dish.

3.7.1.7 By using a large-bore, smooth-tip glass pipette, select mysids from the 

counting dish(es) and place them in the 36 individually numbered 

distribution containers (10-ml beakers are suggested). The mysids are 

assigned two at a time to the 36 containers by using a randomization 

schedule similar to the one presented below. At the end of 

selection/assignment round 1, each container will contain two mysids; at 

the end of round 2, they will contain four mysids; and so on until each 

contains ten mysids.

Table 3-1. Example of a Randomization Schedule

Selection/assignment 

round (2 mysids each)

Place mysid in the numbered distribution containers in the 

random order shown

1 8, 21, 6, 28, 33, 32, 1, 3, 10, 9, 4, 14, 23, 2, 34, 22, 36, 27, 5, 30, 

35, 24, 12, 25, 11, 17, 19, 26, 31, 7, 20, 15, 18, 13, 16, 29.

2 35, 18, 5, 12, 32, 34, 22, 3, 9, 16, 26, 13, 20, 28, 6, 21, 24, 30, 8, 

31, 7, 23, 2, 15, 25, 17, 1, 11, 27, 4, 19, 36, 10, 33, 14, 29.

3 7, 19, 14, 11, 34, 21, 25, 27, 17, 18, 6, 16, 29, 2, 32, 10, 4, 20, 3, 9, 

1, 5, 28, 24, 31, 15, 22, 13, 33, 26, 36, 12, 8, 30, 35, 23.

4 30, 2, 18, 5, 8, 27, 10, 25, 4, 20, 26, 15, 31, 36, 35, 23, 11, 29, 16, 

17, 28, 1, 33, 14, 9, 34, 7, 3, 12, 22, 21, 6, 19, 24, 32, 13.

5 34, 28, 16, 17, 10, 12, 1, 36, 20, 18, 15, 22, 2, 4, 19, 23, 27, 29, 25, 

21, 30, 3, 9, 33, 32, 6, 14, 11, 35, 24, 26, 7, 31, 5, 13, 8.

3.7.1.8 Transfer mysids from the 36 distribution containers to 18 labeled test 

containers in random order. A label is assigned to each of the three 

replicates (A, B, C) of the six test concentrations. Count and record the 96 

hour response in an impartial order.
3-10

3.7.1.9 Repeat Tasks 5–7 (Sections 3.7.1.5 through 3.7.1.7) for each toxicity test. 

A new random schedule should be followed in Tasks 6 and 7 for each test.

Note: If a partial toxicity test is conducted, the procedures described above 

are appropriate and should be used to prepare the single test concentration 

and control, along with the reference toxicant test.

3.7.2 Data Analysis and Interpretation

3.7.2.1 Complete survival data in all test containers at each observation time shall 

be presented in tabular form. If greater than 10% mortality occurs in the 

controls, all data shall be discarded and the experiment repeated. 

Unacceptably high control mortality indicates the presence of important 

stresses on the organisms other than the material being tested, such as 

injury or disease, stressful physical or chemical conditions in the 

containers, or improper handling, acclimation, or feeding. If 10%

mortality or less occurs in the controls, the data may be evaluated and 

reported.

3.7.2.2 A definitive, full bioassay conducted according to the EPA protocol is 

used to estimate the concentration that is lethal to 50% of the test 

organisms that do not die naturally. This toxicity measure is known as the 

median lethal concentration, or LC50. The LC50 is adjusted for natural 

mortality or natural responsiveness. The maximum likelihood estimation 

procedure with the adjustments for natural responsiveness as given by D.J. 

Finney, in Probit Analysis 3rd edition, 1971, Cambridge University Press, 

chapter 7, can be used to obtain the probit model estimate of the LC50 and 

the 95% fiducial (confidence) limits for the LC50. These estimates are 

obtained using the logarithmic transform of the concentration. The 

heterogeneity factor (Finney 1971, pages 70–72) is not used. For a test 

material to pass the toxicity test, according to the requirements stated in 

the offshore oil and gas extraction industry BAT effluent limitations and 

NSPS, the LC50, adjusted for natural responsiveness, must be greater than 

3% suspended particulate phase (SPP) concentration by volume 

unadjusted for the 1 to 9 dilution. Other toxicity test models may be used 

to obtain toxicity estimates provided the modeled mathematical expression 

for the lethality rate must increase continuously with concentration. The 

lethality rate is modeled to increase with concentration to reflect an 

assumed increase in toxicity with concentration even though the observed 

lethality may not increase uniformly because of the unpredictable animal 

response fluctuations.

3.7.2.3 The range finding test is used to establish a reasonable set of test 

concentrations in order to run the definitive test. However, if the lethality 

rate changes rapidly over a narrow range of concentrations, the range 

finding assay may be too coarse to establish an adequate set of test 

concentrations for a definitive test.
3-11

3.7.2.4 The EPA Environmental Research Laboratory in Gulf Breeze, Florida 

prepared a Research and Development Report entitled Acute Toxicity of 

Eight Drilling Fluids to Mysid Shrimp (Mysidopsis bahia), May 1984 

EPA–600/3–84–067. The Gulf Breeze data for drilling fluid number 1 are 

displayed in Table 3-2 for purposes of an example of the probit analysis 

described above. The SAS Probit Procedure (SAS Institute, Statistical 

Analysis System, Cary, North Carolina, 1982) was used to analyze these 

data. The 96-hour LC50 adjusted for the estimated spontaneous mortality

rate is 3.3% SPP with 95% limits of 3.0 and 3.5% SPP with the 1 to 9 

dilution. The estimated spontaneous mortality rate based on all of the data 

is 9.6%.

Table 3-2. Listing of Acute Toxicity Test Data (August 1983 to September 1983) With Eight 

Generic Drilling Fluids and Mysid Shrimp

[fluid N2=1]

Percent Concentration Number Exposed Number Dead (96 hours) Number Alive (96 hours)

0 60 3 57

1 60 11 49

2 60 11 49

3 60 25 35

4 60 48 12

5 60 60 0

3.7.3 The Partial Toxicity Test for Evaluation of Test Material

3.7.3.1 A partial test conducted according to EPA protocol can be used 

economically to demonstrate that a test material passes the toxicity test. 

The partial test cannot be used to estimate the LC50 adjusted for natural 

response.

3.7.3.2 To conduct a partial test follow the test protocol for preparation of the test 

material and organisms. Prepare the control (zero concentration), one test 

concentration (3% suspended particulate phase) and the reference toxicant 

according to the methods of the full test. A range finding test is not used 

for the partial test.

3.7.3.3 Sixty test organisms are used for each test concentration. Find the number 

of test organisms killed in the control (zero percent SPP) concentration in 

the column labeled X0 of Table 3-3. If the number of organisms in the 

control (zero percent SPP) exceeds the table values, then the test is 

unacceptable and must be repeated. If the number of organisms killed in 

the 3% test concentration is less than or equal to corresponding number in 

the column labeled X1 then the test material passes the partial toxicity test. 

Otherwise the test material fails the toxicity test.

3.7.3.4 Data shall be reported as percent suspended particulate phase.
3-12

Table 3-3. Partial Toxicity Test Passing Criteria 

Number Killed in Control (X0) Number Killed in Test (X1)

0 22

1 22

2 23

3 23

4 24

5 24

6 25

3.8 References

1. Borthwick, Patrick W. 1978. Methods for acute static toxicity tests with mysid shrimp 

(Mysidopsis bahia). Bioassay Procedures for the Ocean Disposal Permit Program, [EPA–

600/9–78–010:] March.

2. Nimmo, D.R., T.L. Hamaker, and C.A. Somers. 1978. Culturing the mysid (Mysidopsis 

bahia) in flowing seawater or a static system. Bioassay Procedures for the Ocean 

Disposal Permit Program, [EPA–600/9–78–010]: March.

3. American Public Health Association et al. 1980. Standard Methods for the Examination 

of Water and Wastewater. Washington, DC, 15th Edition: 90–99.

4. U.S. Environmental Protection Agency, September 1991. Methods for Measuring the 

Acute Toxicity of Effluents and Receiving Waters to Freshwater and Marine Organisms. 

EPA/600/4–90/027. Washington, DC, 4th Edition.

5. Finney, D.J. Probit Analysis. Cambridge University Press; 1971.

6. U.S. Environmental Protection Agency, May 1984. Acute Toxicity of Eight Drilling 

Fluids to Mysid Shrimp (Mysidopsis bahia). EPA–600/3–84–067.
4-1

4. PROCEDURE FOR MIXING BASE FLUIDS WITH SEDIMENTS (EPA METHOD 1646)

This procedure describes a method for amending uncontaminated and nontoxic (control) 

sediments with the base fluids that are used to formulate synthetic-based drilling fluids and other 

non-aqueous drilling fluids. Initially, control sediments shall be press-sieved through a 2,000 

micron mesh sieve to remove large debris. Then press-sieve the sediment through a 500 micron 

sieve to remove indigenous organisms that may prey on the test species or otherwise confound 

test results. Homogenize control sediment to limit the effects of settling that may have occurred 

during storage. Sediments should be homogenized before density determinations and addition of 

base fluid to control sediment. Because base fluids are strongly hydrophobic and do not readily 

mix with sediment, care must be taken to ensure base fluids are thoroughly homogenized within 

the sediment. All concentrations are weight-to-weight (mg of base fluid to kg of dry control 

sediment). Sediment and base fluid mixing shall be accomplished by using the following 

method.

4.1 Determining the Wet to Dry Ratio for the Control Sediment

Determine the wet to dry ratio for the control sediment by weighing approximately 10 g 

subsamples of the screened and homogenized wet sediment into tared aluminum weigh pans. Dry

sediment at 105°C for 18–24 h. Remove sediment and cool in a desiccator until a constant 

weight is achieved. Re-weigh the samples to determine the dry weight. Determine the wet/dry

ratio by dividing the net wet weight by the net dry weight:

[4-1]

4.2 Determining the Density of the Wet Control or Dilution Sediment

Determine the density (g/mL) of the wet control or dilution sediment. This shall be used to 

determine total volume of wet sediment needed for the various test treatments.

[4-2]

4.3 Determining the Amount of Base Fluid Needed

To determine the amount of base fluid needed to obtain a test concentration of 500 mg base fluid 

per kg dry sediment use the following formulas:

Determine the amount of wet sediment required:

[4-3]

Determine the amount of dry sediment in kilograms (kg) required for each concentration:

Wet Sediment 

Density (g/mL) ×

Volume of Sediment Required 

per Concentration (mL) =

Weight Wet 

Sediment Required 

per Conc. (g)

Mean Wet Sediment Weight (g) = Wet Sediment Density (g/mL) Mean Wet Sediment Volume (mL)

Wet Sediment Weight (g) = Wet to Dry Ratio Dry Sediment Weight (g)
4-2

[4-4]

Finally, determine the amount of base fluid required to spike the control sediment at each 

concentration:

[4-5]

For spiking test substances other than pure base fluids (e.g., whole mud formulations), determine 

the spike amount as follows:

[4-6]

4.4 Primary Mixing

For primary mixing, place appropriate amounts of weighed base fluid into stainless mixing 

bowls, tare the vessel weight, then add sediment and mix with a high-shear dispersing impeller 

for 9 minutes. The concentration of base fluid in sediment from this mix, rather than the nominal 

concentration, shall be used in calculating LC50 values.

4.5 Testing for Homogeneity of Base Fluid

Tests for homogeneity of base fluid in sediment are to be performed during the procedure 

development phase. Because of difficulty of homogeneously mixing base fluid with sediment, it 

is important to demonstrate that the base fluid is evenly mixed with sediment. The sediment shall 

be analyzed for total petroleum hydrocarbons (TPH) using EPA Methods 3550A and 8015M, 

with samples taken both prior to and after distribution to replicate test containers. Base-fluid 

content is measured as TPH. After mixing the sediment, a minimum of three replicate sediment 

samples shall be taken prior to distribution into test containers. After the test sediment is 

distributed to test containers, an additional three sediment samples shall be taken from three test 

containers to ensure proper distribution of base fluid within test containers. Base-fluid content 

results shall be reported within 48 hours of mixing. The coefficient of variation (CV) for the 

replicate samples must be less than 20%. If base-fluid content results are not within the 20% CV 

limit, the test sediment shall be remixed. Tests shall not begin until the CV is determined to be 

below the maximum limit of 20%. During the test, a minimum of three replicate containers shall 

be sampled to determine base-fluid content during each sampling period.

4.6 Commencing the Sediment Toxicity Test

Mix enough sediment in this way to allow for its use in the preparation of all test concentrations 

and as a negative control. When commencing the sediment toxicity test, range-finding tests may

be required to determine the concentrations that produce a toxic effect if these data are otherwise 

unavailable. The definitive test shall bracket the LC50, which is the desired endpoint. The results 

for the base fluids shall be reported in mg of base fluid per kg of dry sediment.

Conc. Desired 

(mg/kg)

×

Dry Weight 

Sediment (kg) = Base Fluid Required (mg)

Conc. Desired 

(mL/kg)

×

Dry Weight 

Sediment (kg)

×

Test Substance 

Density (g/mL) =

Test Substance 

Required (g)

Wet Sediment per 

Concentration (g) × 1 kg = Dry Weight Sediment (kg)

Mean Wet to Dry Ratio 1,000 g
4-3

4.7 References

1. American Society for Testing and Materials (ASTM). 1996. Standard Guide for 

Collection, Storage, Characterization, and Manipulation of Sediments for Toxicological 

Testing. ASTM E 1391–94. Annual Book of ASTM Standards, Volume 11.05, pp. 805–

825.

2. Ditsworth, G.R., D.W. Schults and J.K.P. Jones. 1990. Preparation of benthic substrates 

for sediment toxicity testing, Environ. Toxicol. Chem. 9:1523–1529.

3. Suedel, B.C., J.H. Rodgers, Jr. and P.A. Clifford. 1993. Bioavailability of fluoranthene in

freshwater sediment toxicity tests. Environ. Toxicol. Chem. 12:155–165.

4. U.S. EPA. 1994. Methods for Assessing the Toxicity of Sediment-associated 

Contaminants with Estuarine and Marine Amphipods. EPA/600/R–94/025. Office of 

Research and Development, Washington, DC.

5. U.S. EPA. 2001. Effluent Limitations Guidelines and New Source Performance 

Standards for the Oil and Gas Extraction Point Source Category. Federal Register, 66: 

6849 (22 January 2001). 

6. U.S. EPA. 2001. Effluent Limitations Guidelines and New Source Performance 

Standards for the Oil and Gas Extraction Point Source Category: Correction. Federal 

Register, 66: 30811 (8 June 2001).
5-1

5. PROTOCOL FOR THE DETERMINATION OF DEGRADATION OF NON-AQUEOUS BASE 

FLUIDS IN A MARINE CLOSED BOTTLE BIODEGRADATION TEST SYSTEM: MODIFIED 

ISO 11734:1995 (EPA METHOD 1647)

5.1 Summary of Method

This method determines the anaerobic degradation potential of mineral oils, paraffin oils and 

non-aqueous fluids (NAF) in sediments. These substrates are base fluids for formulating offshore 

drilling fluids. The test evaluates base fluid biodegradation rates by monitoring gas production 

due to microbial degradation of the test fluid in natural marine sediment. 

The test procedure places a mixture of marine/estuarine sediment, test substrate (hydrocarbon or 

controls) and seawater into clean 120 ml (150 ml actual volume) Wheaton serum bottles. The test 

is run using four replicate serum bottles containing 2,000 mg carbon/kg dry weight concentration 

of test substrate in sediment. The use of resazurin dye solution (1 ppm) evaluates the anaerobic 

(redox) condition of the bottles (dye is blue when oxygen is present, reddish in low oxygen 

conditions and colorless if oxygen free). After capping the bottles, a nitrogen sparge removes air 

in the headspace before incubation begins. During the incubation period, the sample should be 

kept at a constant temperature of 29 ±1°C. Gas production and composition is measured 

approximately every two weeks. The samples need to be brought to ambient temperature before 

making the measurements. Measure gas production using a pressure gauge. Barometric pressure 

is measured at the time of testing to make necessary volume adjustments. 

ISO 11734:1995 specifies that total gas is the standard measure of biodegradation. While 

modifying this test for evaluating biodegradation of NAFs, methane was also monitored and 

found to be an acceptable method of evaluating biodegradation. Section 5.7 contains the 

procedures used to follow biodegradation by methane production. Measurement of either total 

gas or methane production is permitted. If methane is followed, determine the composition of the 

gas by using gas chromatography (GC) analysis at each sampling. At the end of the test when gas 

production stops, or at around 275 days, an analysis of sediment for substrate content is possible. 

Common methods which have been successfully used for analyzing NAFs from sediments are 

listed in Section 5.8.

5.2 System Requirements

This environmental test system has three phases, spiked sediment, overlying seawater, and a gas 

headspace. The sediment/test compound mixture is combined with synthetic sea water and 

transferred into 120 mL serum bottles. The total volume of sediment/sea water mixture in the 

bottles is 75 mL. The volume of the sediment layer will be approximately 50 mL, but the exact 

volume of the sediment will depend on sediment characteristics (wet:dry ratio and density). The 

amount of synthetic sea water will be calculated to bring the total volume in the bottles to 75 mL. 

The test systems are maintained at a temperature of 29 ±1

oC during incubation. The test systems 

are brought to ambient temperatures prior to measuring pressure or gas volume. 

5.2.1 Sample Requirements

The concentration of base fluids are at least 2,000 mg carbon test material/kg dry

sediment. Carbon concentration is determined by theoretical composition based on 

the chemical formula or by chemical analysis by ASTM D5291-96. Sediments with 
5-2

positive, intermediate and negative control substances as well as a C16 –C18 internal 

olefin type base fluid will be run in conjunction with test materials under the same 

conditions. The positive control is ethyl oleate (CAS 111-62-6), the intermediate 

control is 1-hexadecene (CAS 629-73-2), and the negative control is squalane (CAS 

111-01-3). Controls must be of analytical grade or the highest grade available. Each 

test control concentration should be prepared according to the mixing procedure

described in Section 5.3.1.

Product names will be used for examples or clarification in the following text. Any

use of trade or product names in this publication is for descriptive use only, and dos 

not constitute endorsement by EPA or the authors

5.2.2 Seawater Requirements

Synthetic seawater at a salinity of 25 ±1 ppt should be used for the test. The synthetic

seawater should be prepared by mixing a commercially available artificial seawater 

mix, into high purity distilled or de-ionized water. The seawater should be aerated 

and allowed to age for approximately one month prior to use.

5.2.3 Sediment Requirements

The dilution sediment must be from a natural estuarine or marine environment and be

free of the compounds of interest. The collection location, date and time will be 

documented and reported. The sediment is prepared by press-sieving through a 2,000-

micron mesh sieve to remove large debris, then press-sieving through a 500-micron 

sieve to remove indigenous organisms that may confound test results. The water 

content of the sediment should be less than 60% (w/w) or a wet to dry ratio of 2.5. 

The sediment should have a minimum organic matter content of 3% (w/w) as 

determined by ASTM D2974-87 (95) (Method A and D and calculate organic matter 

as in Section 12 of method ASTM D2974-87). 

To reduce the osmotic shock to the microorganisms in the sediment the salinity of the

sediment’s pore water should be between 20-30 ppt. Sediment should be used for 

testing as soon as possible after field collection. If required, sediment can be stored in 

the dark at 4°C with 3-6 inches of overlying water in a sealed container for a 

maximum period of 2 months prior to use. 

5.3 Test Set Up

The test is set up by first mixing the test or control substrates into the sediment inoculum, then 

mixing in seawater to make a pourable slurry. The slurry is then poured into serum bottles, which 

are then flushed with nitrogen and sealed.

5.3.1 Mixing Procedure

Because base fluids are strongly hydrophobic and do not readily mix with sediments, 

care must be taken to ensure base fluids are thoroughly homogenized within the 

sediment. All concentrations are weight-to-weight comparisons (mg of base fluid to 
5-3

kg of dry control sediment). Sediment and base fluid mixing will be accomplished by

using the following method.

5.3.1.1 Determine the wet to dry weight ratio for the control sediment by

weighing approximately 10 sub-samples of approximately 1 g each of the 

screened and homogenized wet sediment into tared aluminum weigh pans. 

Dry sediment at 105°C for18-24 h. Remove the dried sediments and cool 

in a desiccator. Repeat the drying, cooling, and weighing cycle until a 

constant weight is achieved (within 4% of previous weight). Re-weigh the 

samples to determine the dry weight. Calculate the mean wet and dry

weights of the 10 sub samples and determine the wet/dry ratio by dividing 

the mean wet weight by the mean dry weight using Equation 5-1. This is 

required to determine the weight of wet sediment needed to prepare the 

test samples.

[5-1]

5.3.1.2 Determine the density (g/ml) of the wet sediment. This will be used to 

determine total volume of wet sediment needed for the various test 

treatments. One method is to tare a 5 ml graduated cylinder and add about 

5 ml of homogenized sediment. Carefully record the volume then weigh 

this volume of sediment. Repeat this a total of three times. To determine 

the wet sediment density, divide the weight by volume per the following

formula:

[5-2]

5.3.1.3 Determine the amount of base fluid to be spiked into wet sediment in 

order to obtain the desired initial base fluid concentration of 2,000 mg 

carbon/kg dry weight. An amount of wet sediment that is the equivalent of 

30 g of dry sediment will be added to each bottle. A typical procedure is 

to prepare enough sediment for 8 serum bottles (3 bottles to be sacrificed 

at the start of the test, 4 bottles incubated for headspace analysis, and 

enough extra sediment for 2 extra bottles). Extra sediment is needed 

because some of the sediment will remain coated onto the mixing bowl 

and utensils. Experience with this test may indicate that preparing larger 

volumes of spiked sediment is a useful practice, then the following 

calculations should be adjusted accordingly.

a. Determine the total weight of dry sediment needed to add 30 g dry

sediment to 8 bottles. If more bottles are used then the calculations 

should be modified accordingly. For example:

30 g dry sediment per bottle × 8 = 240 g dry sediment [5-3]

Mean Wet Sediment Weight (g)

= Wet Sediment Density (g/mL) Mean Wet Sediment Volume (mL)

Mean Wet Sediment Weight (g)

= Wet to Dry Ratio Mean Dry Sediment Weight (g)
5-4

b. Determine the weight of base fluid, in terms of carbon, needed to 

obtain a final base fluid concentration of 2,000 mg carbon/kg dry

weight. For example:

[5-4]

c. Convert from mg of carbon to mg of base fluid. This calculation will 

depend on the % fraction of carbon present in the molecular structure 

of each base fluid. For the control fluids, ethyl oleate is composed of

77.3% carbon, hexadecene is composed of 85.7% carbon, and 

squalane is composed of 85.3% carbon. The carbon fraction of each 

base fluid should be supplied by the manufacturer or determined 

before use. ASTM D5291-96 or equivalent will used to determine 

composition of fluid.

To calculate the amount of base fluid to add to the sediment, divide the 

amount of carbon (480 mg) by the percent fraction of carbon in the 

fluid.

For example, the amount of ethyl oleate added to 240 g dry weight 

sediment can be calculated from the following equation:

[5-5]

Therefore, add 621 mg of ethyl oleate to 240 g dry weight sediment for 

a final concentration of 2,000 mg carbon/kg sediment dry weight.

5.3.1.4 Mix the calculated amount of base fluid with the appropriate weight of wet 

sediment.

a. Use the wet:dry ratio to convert from g sediment dry weight to g 

sediment wet weight, as follows:

240 g dry sediment ×wet:dry ratio = g wet sediment needed [5-6]

b. Weigh the appropriate amount of base fluid (calculated in Section 

5.3.1.3c) into stainless mixing bowls, tare the vessel weight, then add 

the wet sediment calculated in Equation 5-5, and mix with a high shear 

dispersing impeller for 9 minutes.

The sediment is now mixed with synthetic sea water to form a slurry

that will be transferred into the bottles. 

5.3.2 Creating Seawater/Sediment Slurry

Given that the total volume of sediment/sea water slurry in each bottle is to be 75 mL, 

determine the volume of sea water to add to the wet sediment.

480 mg carbon

= 621 mg ethyl oleate (77.3 ÷ 100)

2,000 mg carbon

×

240 g

= 480 mg carbon Per kg dry sediment 1,000
5-5

5.3.2.1 If each bottle is to contain 30 g dry sediment, calculate the weight, and 

then the volume, of wet sediment to be added to each bottle

30 g dry sediment × wet:dry ratio = g wet sediment added to each bottle [5-7]

[5-8]

5.3.2.2 Calculate volume of sea water to be added to each bottle

75 mL total volume – mL wet sediment (from Eq. 8) = mL of sea water [5-9]

5.3.2.3 Determine the ratio of sea water to wet sediment (volume:volume) in each 

bottle

[5-10]

5.3.2.4 Convert the wet sediment weight from Equation 5-6 into a volume using 

the sediment density.

g wet sediment (Eq. 5-6) density = volume (mL) of sediment [5-11]

5.3.2.5 Determine the amount of sea water to mix with the wet sediment.

[5-12]

Mix sea water thoroughly with wet sediment to form a sediment/sea water 

slurry.

5.3.3 Bottling the Sediment Seawater Slurry

The total volume of sediment/sea water slurry in each bottle is to be 75 mL. Convert 

the volume (mL) of sediment/sea water slurry into a weight (g) using the density of 

the sediment and the seawater.

5.3.3.1 Determine the weight of sediment to be added to each bottle

mL sediment (Eq. 5-8) × density of wet sediment (g/mL) = g wet sediment [5-13]

5.3.3.2 Determine the weight of sea water to be added to each bottle

mL sea water (Eq. 5-9) × density of sea water (1.01 g/mL) = g sea water [5-14]

5.3.3.3 Determine weight of sediment/sea water slurry to be added to each bottle

g wet sediment (Eq. 5-13) + g sea water (Eq. 5-14) = g sediment/sea water slurry [5-15]

This should provide each bottle with 30 g dry sediment in a total volume of 75 mL.

mL wet sediment 

(Eq. 5-11)

×

Sea water:sediment ratio 

(Eq. 5-10) = mL sea water to add to wet sediment

g wet sediment = mL wet sediment Density (g/mL) of wet sediment

Volume sea water per bottle (Eq. 5-9) = Ratio of sea water:wet sediment Volume sediment water per bottle (Eq. 5-8)
5-6

5.3.3.4 Putting the sediment:seawater slurry in the serum bottles.

Note: The slurry will need to be constantly stirred to keep the sediment 

suspended.

Place a tared serum bottle on a balance and add the appropriate amount of 

slurry to the bottle using a funnel. Once the required slurry is in the bottle 

remove the funnel, add 2-3 drops (25 μL) of a 1 gram/L resazurin dye 

stock solution. Cap the bottle with a butyl rubber stopper (Bellco Glass, 

Part #2048- 11800) and crimp with an aluminum seal (Bellco Glass Part 

#2048-11020).

Using a plastic tube with a (23 gauge, 1 inch long) needle attached to one 

side and a nitrogen source to the other, puncture the serum cap with the 

needle. Puncture the serum cap again with a second needle to sparge the 

bottle’s headspace of residual air for two minutes. The nitrogen should be 

flowing at no more than 100 mL/min to encourage gentle displacement of 

oxygenated air with nitrogen. Faster nitrogen flow rates would cause 

mixing and complete oxygen removal would take much longer. Remove 

the nitrogen needle first to avoid any initial pressure problems. The second 

(vent) needle should be removed within 30 seconds of removing the 

nitrogen needle.

Triplicate blank test systems are prepared, with similar quantities of 

sediment and seawater without any base fluid. Incubate in the dark at a 

constant temperature of 29 1°C.

Record the test temperature. The test duration is dependent on base fluid 

performance, but at a maximum should be no more than 275 days. Stop 

the test after all base fluids have achieved a plateau of gas production. At 

termination, base fluid concentrations can be verified in the terminated 

samples by extraction and GC analysis according to Section 5.8.

5.4 Concentration Verification Chemical Analyses

Because of the difficulty of homogeneously mixing base fluid with sediment, it is important to 

demonstrate that the base fluid is evenly mixed within the sediment sea water slurry that was 

added to each bottle. Of the seven serum bottles set up for each test or control condition, three 

are randomly selected for concentration verification analyses. These should be immediately

placed at 4C and a sample of sediment from each bottle should be analyzed for base fluid 

content as soon as possible. The coefficient of variation (CV) for the replicate samples must be 

less than 20%. The results should show recovery of at least 70% of the spiked base fluid. Use an 

appropriate analytical procedure described in Section 5.8 to perform the extractions and analyses. 

If any set of sediments fail the criteria for concentration verification, then the corrective action 

for that set of sediments is also outlined in Section 5.8.

The nominal concentrations and the measured concentrations from the three bottles selected for 

concentration verification should be reported for the initial test concentrations. The coefficient of 

variation (CV) for the replicate samples must be less than 20%. If base fluid content results are 
5-7

not within the 20% CV limit, the test must be stopped and restarted with adequately mixed 

sediment.

5.5 Gas Monitoring Procedures

Biodegradation is measured by total gas as specified in ISO 11734:1995. Methane 

production can also

be tracked and is described in Section 5.7.

5.5.1 Total Gas Monitoring Procedures

Bottles should be brought to room temperature before readings are taken. The bottles 

are observed to confirm that the resazurin has not oxidized to pink or blue. Total gas 

production in the culture bottles should be measured using a pressure transducer (one 

source is Biotech International). The pressure readings from test and control cultures 

are evaluated against a calibration curve created by analyzing the pressure created by

known additions of gas to bottles established identically to the culture bottles. Bottles 

used for the standard curve contain 75 mL of water, and are sealed with the same 

rubber septa and crimp cap seals used for the bottles containing sediment. After the 

bottles used in the standard curve have been sealed, a syringe needle inserted through 

the septa is used to equilibrate the pressure inside the bottles to the outside 

atmosphere. The syringe needle is removed and known volumes of air are injected 

into the headspace of the bottles. Pressure readings provide a standard curve relating 

the volume of gas injected into the bottles and headspace pressure. No less than three 

points may be used to generate the standard curve. A typical standard curve may use 

0, 1, 5, 10, 20 and 40 ml of gas added to the standard curve bottles.

The room temperature and barometric pressure (to two digits) should be recorded at 

the time of sampling. One option for the barometer is Fisher Part #02-400 or 02-401. 

Gas production by the sediment is expressed in terms of the volume (mL) of gas at 

standard temperature (0°C = 273°K) and pressure (1 atm = 30 inches of Hg) using Eq. 

5-16.

[5-16]

Where: 

V2 = Volume of gas production at standard temperature and pressure

P1 = Barometric pressure on day of sampling (inches of Hg)

V1 = Volume of gas measured on day of sampling (mL)

T2 = Standard temperature = 273°K

T1 = Temperature on day of sampling (°C + 273 = °K)

P2 = Standard pressure = 30 inches Hg

An estimate can be made of the total volume of anaerobic gas that will be produced in 

the bottles. The gas production measured for each base fluid can be expressed as a 

percent of predicted total anaerobic gas production.

V2 =

P1 × V1 × T2

T1 × P2
5-8

5.5.1.1 Calculate the total amount of carbon in the form of the base fluid present 

in each bottle

Each bottle is to contain 30 g dry weight sediment. The base fluid 

concentration is 2,000 mg carbon/kg dry weight sediment. Therefore:

2,000 mg carbon/kg sediment × (30 g ÷ 1,000) = 60 mg carbon per bottle [5-17]

5.5.1.2 Theory states that anaerobic microorganisms will convert 1 mole of 

carbon substrate into 1 mole of total anaerobic gas production

Calculate the number of moles of carbon in each bottle.

The molecular weight of carbon is 12 (i.e., 1 mole of carbon = 12 g). 

Therefore, the number of moles of carbon in each bottle can be 

calculated.

[5-18]

5.5.1.3 Calculate the predicted volume of anaerobic gas

One mole of gas equals 22.4 L (at standard temperature and pressure), 

therefore, 

0.005 moles × 22.4 L = 0.112L (or 112 mL total gas production) [5-19]

5.5.2 Gas Venting

If the pressure in the serum bottle is too great for the pressure transducer or syringe, 

some of the excess gas must be wasted. The best method to do this is to vent the 

excess gas right after measurement. To do this, remove the barrel from a 10-mL 

syringe and fill it 1/3 full with water. This is then inserted into the bottle 

through the stopper using a small diameter (high gauge) needle. The excess 

pressure is allowed to vent through the water until the bubbles stop. This allows 

equalization of the pressure inside the bottle to atmospheric without 

introducing oxygen. The amount of gas vented (which is equal to the volume 

determined that day) must be kept track of each time the bottles are vented. A 

simple way to do this in a spreadsheet format is to have a separate column in 

which cumulative vented gas is tabulated. Each time the volume of gas in the 

cultures is analyzed, the total gas produced is equal to the gas in the culture at 

that time plus the total of the vented gas.

To keep track of the methane lost in the venting procedure, multiply the amount of 

gas vented each time by the corrected % methane determined on that day. The answer 

gives the volume of methane wasted. This must be added into the cumulative totals 

similarly to the total gas additions.

60 mg carbon per bottle/1,000 = 0.005 moles carbon 12 g/mole
5-9

5.6 Test Acceptability and Interpretation

5.6.1 Test Acceptability

At day 275 or when gas production has plateaued, whichever is first, the controls are 

evaluated to confirm that the test has been performed appropriately. In order for this 

modification of the closed bottle biodegradation test to be considered acceptable, all 

the controls must meet the biodegradation levels indicated in Table 5-1. The 

intermediate control hexadecene must produce at least 30% of the theoretical gas 

production. This level may be reexamined after two years and more data has been 

generated.

Table 5-1. Test Acceptability Criteria

Concentration Percent Biodegradability as a Function of Gas Measurement

Positive control Squalane negative control Hexadecene intermediate control

2,000 mg carbon/kg ≥60% theoretical ≤5% theoretical ≥30% theoretical

5.6.2 Interpretation

In order for a fluid to pass the closed bottle test, the biodegradation of the base fluid 

as indicated by the total amount of total gas (or methane) generated once gas 

production has plateaued (or at the end of 275 days, which ever is first) must be 

greater than or equal to the volume of gas (or methane) produced by the reference 

standard (internal elefin or ester).

The method for evaluating the data to determine whether a fluid has passed the 

biodegradation test must use the equations:

[5-20]

Where:

NAF = Stock base fluid being tested for compliance

Reference fluid = C16 –C18 internal olefin or C12 –C14 or C8 ester reference

fluid

5.7 Methane Measurement

5.7.1 Methane Monitoring Procedures

The use of total gas production alone may result in an underestimation of the actual 

metabolism occurring since CO2 is slightly soluble in water. An acceptable alternative 

method is to monitor methane production and total gas production. This is easily done 

using GC analysis. A direct injection of headspace gases can be made into a GC using 

almost any packed or capillary column with an FID detector. Unless volatile fuels or 

solvents are present in the test material or the inocula, the only component of the 

headspace gas that can be detected using an FID detector is methane. The percent 

% Theoretical gas production of reference fluid ≤ 1.0 % Theoretical gas production of NAF
5-10

methane in the headspace gas is determined by comparing the response of the sample 

injections to the response from injections of known percent methane standards. The 

percent methane is corrected for water vapor saturation using Eq. 5-21 and then 

converted to a volume of dry methane using Eq. 5-22.

[5-21]

Where:

D = The density of water vapor at saturation (g/m3, can be found in 

CRC Handbook of Chemistry and Physics) for the temperature 

of sampling.

[5-22]

Where: 

VCH4 = The volume of methane in the bottle

S = Volume of excess gas production (measured with a pressure 

transducer)

V = Volume of the headspace in the culture bottle (total volume - liquid 

phase)

P = Barometric pressure (mm Hg, measured with barometer)

T = Temperature (C)

Pw = Vapor pressure of water at T (mm Hg, can be found in CRC 

Handbook of Chemistry and Physics)

CH4 = % methane in headspace gas (after correction for water vapor)

The total volume of serum bottles sold as 125 mL bottles (Wheaton) is 154.8 mL.

The volumes of methane produced are then compared to the volumes of methane in 

the controls to determine if a significant inhibition of methane production or a 

significant increase of methane production has been observed. Effective statistical 

analyses are important, as variability in the results is common due to the 

heterogeneity of the inoculum’s source. It is also common to observe that the timing 

of the initiation of culture activity is not equal in all of the cultures. Expect a great 

variability over the period when the cultures are active, some replicates will start 

sooner than others, but all of the replicates should eventually reach similar levels of 

base fluid degradation and methane production.

5.7.2 Expected Methane Production Calculations

The amount of methane expected can be calculated using the equation of Symons and 

Buswell (Eq. 5-23). In the case of complete mineralization, all of the carbon will 

appear as wither CO2 or CH4, thus the total moles of gas produced will be equal to the 

total moles of carbon in the parent molecule. The use of the Buswell equation allows 

you to calculate the effects the redox potential will have on the distribution of the 

VCH4 (ml) = (S + V) × P – Pw×

CH4

×

273

T + 273 100 760

Corrected % CH4 =

% CH4

1 –

D × 22.4 L/mol

18 g/mol × 1,000
5-11

products in methanogenic cultures. More reduced electron donors will allow the 

production of more methane, while more oxidized electron donors will cause a 

production of more carbon dioxide.

CnHaObNcSd + (n-a/4 -b/2 + 7c/4 + d/2) H2O → (n/2 -a/8+b/4-5c/8 + d/4) CO2 + 

(n/2 +a/8 -b/4 -3c/8-d/4) CH4 + cNH4HCO3 + dH2S [5-23]

An example calculation of the expected methane volume in a culture fed 2,000 mg/kg

hexadecene is as follows. The application of Symons and Buswell’s equation reveals

that hexadecene (C16H32) will yield 4 moles of CO2 and 12 moles of CH4. Assuming 

30 g of dry sediment are added to the bottles with 2,334 mg hexadecene/kg dry

sediment (i.e., equivalent to 2,000 mg carbon/kg dry sediment) the calculation is as 

follows.

[5-24]

By subtracting the average amount of methane in control bottles from the test bottles

and then dividing by the expected volume an evaluation of the completion of the

process may be conducted.

5.8 Concentration Verification Analysis

The Concentration Verification analysis is required at the beginning of the test to ensure 

homogeneity and confirm that the required amount of fluid was delivered to the sediments at the 

start of the test.

5.8.1 Three samples per fluid need to be analyzed and achieve ≤20% Coefficient of 

Variability and an average of ≥70% to ≤120% of fluid delivered to sediment.

5.8.2 If a third party performs the analysis, then the laboratory should be capable of 

delivering the homogeneity data within seven days, in order to identify any samples 

that do not meet the homogeneity requirement as quickly as possible.

5.8.3 If one sediment/fluid set, out a multiple set batch of samples, fails these criteria, then 

that one set of samples must be discarded and a fresh set of spiked sediment prepared, 

started, and analyzed to ensure homogeneity. The same stock sediment is used to 

prepare the replacement set(s). The remaining sets do not need to be re-mixed or 

restarted.

5.8.4 The re-mixed set(s) will need to be run the additional days as appropriate to ensure 

that the total number of days is the same for all sets of bottles, even though the 

specific days are not aligned.

5.8.5 Re-mixing of bottle sets can be performed multiple times as a result of a failure of the

analytical criteria, until the holding time for the stock sediment has expired (60 days). 

If the problem set(s) has not fallen within the acceptable analytical criteria by then, it 

12 mole CH4

×

22.4 L

×

1,000 

ml

×

1 mole 

hexadecene

×

23 g 

hexadecene

×

0.03 kg = 84 (ml) mole 

hexadecene

mole CH4 L 224.4 g 

hexadecene

kg dry soil culture
5-12

must not be part of the batch of bottles run. If the problem batch is one of the 

controls, and those controls were not successfully prepared when the sediment 

holding time expired, then the entire test must be restarted.

5.9 Program Quality Assurance and Quality Control

5.9.1 Calibration

5.9.1.1 All equipment / instrumentation will be calibrated in accordance with the 

test method or the manufacture's instructions and may be scheduled or 

triggered.

5.9.1.2 Where possible, standards used in calibration will be traceable to a 

nationally recognized standard (e.g., certified standard by NIST).

5.9.1.3 All calibration activities will be documented and the records retained.

5.9.1.4 The source, lot, batch number, and expiration date of all reagents used 

with be documented and retained.

5.9.2 Maintenance

5.9.2.1 All equipment / instrumentation will be maintained in accordance with the 

test method or the manufacture's instructions and may be scheduled or 

triggered.

5.9.2.2 All maintenance activities will be documented and the records retained.

5.9.3 Data Management and Handling

5.9.3.1 All primary (raw) data will be correct, complete, without selective 

reporting, and will be maintained.

5.9.3.2 Hand-written data will be recorded in lab notebooks or electronically at 

the time of observation.

5.9.3.3 All hand-written records will be legible and amenable to reproduction by

electrostatic copiers.

5.9.3.4 All changes to data or other records will be made by:

a. Using a single line to mark-through the erroneous entry (maintaining 

original data legibility)

b. Write the revision

c. Initial, date, and provide revision code (see attached or laboratory’s 

equivalent)
5-13

5.9.3.5 All data entry, transcriptions, and calculations will be verified by a 

qualified person.

a. Verification will be documented by initials of verifier and date

5.9.3.6 Procedures will be in place to address data management procedures used 

(at minimum):

a. Significant figures

b. Rounding practices

c. Identification of outliers in data series

d. Required statistics

5.9.4 Document Control

5.9.4.1 All technical procedures, methods, work instructions, standard operating 

procedures must be documented and approved by laboratory management 

prior to the implementation.

5.9.4.2 All primary data will be maintained by the contractor for a minimum of 

five (5) years.

5.9.5 Personnel and Training

5.9.5.1 Only qualified personnel shall perform laboratory activities.

5.9.5.2 Records of staff training and experience will be available. This will 

include initial and refresher training (as appropriate).

5.9.6 Test Performance

5.9.6.1 All testing will done in accordance with the specified test methods.

5.9.6.2 Receipt, arrival condition, storage conditions, dispersal, and accountability

of the test article will be documented and maintained.

5.9.6.3 Receipt or production, arrival or initial condition, storage conditions, 

dispersal, and accountability of the test matrix (e.g., sediment or artificial 

seawater) will be documented and maintained.

5.9.6.4 Source, receipt, arrival condition, storage conditions, dispersal, and 

accountability of the test organisms (including inoculum) will be 

documented and maintained.

5.9.6.5 Actual concentrations administered at each treatment level will be verified 

by appropriate methodologies.
5-14

5.9.6.6 Any data originating at a different laboratory will be identified and the 

laboratory fully referenced in the final report.

5.9.7 The following references identify analytical methods that have historically been 

successful for achieving the analytical quality criteria.

5.9.7.1 Continental Shelf Associates report 1998. Joint EPA/Industry screening 

survey to assess the deposition of drill cuttings and associated synthetic 

based mud on the seabed of the Louisiana continental shelf, Gulf of 

Mexico. Analysis by Charlie Henry report Number IES/RCAT97-36 GCFID and GC/MS.

5.9.7.2 EPA Method 3550 for extraction with EPA Method 8015 for GC-FID.

5.9.7.3 Webster, L; Mackie, P.R.; Hird, S.J.; Munro, P.D.; Brown, N.A. and 

Moffatt, C.F. (1997) Development of Analytical Methods for the 

Determination of Synthetic Mud Base Fluids in Marine Sediments Analyst 

122:1485-1490.

5.9.7.4 Munro, P.D., B Croce, C.F. Moffet, N.A Brown, A.D. McIntosh, S.J. 

Hird, R.M. Stagg. 1998. Solid-phase test for comparison for degradation 

rates of synthetic mud base fluids used in the off shore drilling industry. 

Environ. Toxicol. Chem. 17:1951-1959.
6-1

6. DETERMINATION OF CRUDE OIL CONTAMINATION IN NON-AQUEOUS DRILLING FLUIDS 

BY GAS CHROMATOGRAPHY/MASS SPECTROMETRY (GC/MS) (EPA METHOD 1655)

6.1 Scope and Application

6.1.1 This method determines crude (formation) oil contamination, or other petroleum oil 

contamination, in non-aqueous drilling fluids (NAFs) by comparing the gas

chromatography/mass spectrometry (GC/MS) fingerprint scan and extracted ion scans 

of the test sample to that of an uncontaminated sample.

6.1.2 This method can be used for monitoring oil contamination of NAFs or monitoring oil 

contamination of the base fluid used in the NAF formulations.

6.1.3 Any modification of this method beyond those expressly permitted shall be 

considered as a major modification subject to application and approval of alternative 

test procedures under 40 CFR 136.4 and 136.5.

6.1.4 The gas chromatography/mass spectrometry portions of this method are restricted to 

use by, or under the supervision of analysts experienced in the use of GC/MS and in 

the interpretation of gas chromatograms and extracted ion scans. Each laboratory that 

uses this method must generate acceptable results using the procedures described in 

Sections 6.7, 6.9.2, and 6.12.

6.2 Summary of Method

6.2.1 Analysis of NAF for crude oil contamination is a step-wise process. The analyst first 

performs a qualitative assessment of the presence or absence of crude oil in the 

sample. If crude oil is detected during this qualitative assessment, the analyst must 

perform a quantitative analysis of the crude oil concentration.

6.2.2 A sample of NAF is centrifuged to obtain a solids free supernate.

6.2.3 The test sample is prepared by removing an aliquot of the solids free supernate, 

spiking it with internal standard, and analyzing it using GC/MS techniques. The 

components are separated by the gas chromatograph and detected by the mass 

spectrometer.

6.2.4 Qualitative identification of crude oil contamination is performed by comparing the 

Total Ion Chromatograph (TIC) scans and Extracted Ion Profile (EIP) scans of test 

sample to that of uncontaminated base fluids, and examining the profiles for 

chromatographic signatures diagnostic of oil contamination.

6.2.5 The presence or absence of crude oil contamination observed in the full scan profiles 

and selected extracted ion profiles determines further sample quantitation and 

reporting requirements.

6.2.6 If crude oil is detected in the qualitative analysis, quantitative analysis must be 

performed by calibrating the GC/MS using a designated NAF spiked with known 

concentrations of a designated oil.
6-2

6.2.7 Quality is assured through reproducible calibration and testing of GC/MS system and 

through analysis of quality control samples.

6.3 Definitions

6.3.1 A NAF is one in which the continuous phase is a water immiscible fluid such as an 

oleaginous material (e.g., mineral oil, enhance mineral oil, paraffinic oil, or synthetic 

material such as olefins and vegetable esters).

6.3.2 TIC—Total Ion Chromatograph.

6.3.3 EIP—Extracted Ion Profile.

6.3.4 TCB—1,3,5-trichlorobenzene is used as the internal standard in this method.

6.3.5 SPTM—System Performance Test Mix standards are used to establish retention times 

and monitor detection levels.

6.4 Interferences and Limitations

6.4.1 Solvents, reagents, glassware, and other sample processing hardware may yield 

artifacts and/or elevated baselines causing misinterpretation of chromatograms.

6.4.2 All Materials used in the analysis shall be demonstrated to be free from interferences 

by running method blanks. Specific selection of reagents and purification of solvents 

by distillation in all-glass systems may be required.

6.4.3 Glassware shall be cleaned by rinsing with solvent and baking at 400°C for a 

minimum of 1 hour.

6.4.4 Interferences may vary from source to source, depending on the diversity of the 

samples being tested.

6.4.5 Variations in and additions of base fluids and/or drilling fluid additives (emulsifiers, 

dispersants, fluid loss control agents, etc.) might also cause interferences and 

misinterpretation of chromatograms.

6.4.6 Difference in light crude oils, medium crude oils, and heavy crude oils will result in 

different responses and thus different interpretation of scans and calculated 

percentages.

6.5 Safety

6.5.1 The toxicity or carcinogenicity of each reagent used in this method has not been 

precisely determined; however each chemical shall be treated as a potential health 

hazard. Exposure to these chemicals should be reduced to the lowest possible level.

6.5.2 Unknown samples may contain high concentration of volatile toxic compounds. 

Sample containers should be opened in a hood and handled with gloves to prevent 
6-3

exposure. In addition, all sample preparation should be conducted in a fume hood to 

limit the potential exposure to harmful contaminates.

6.5.3 This method does not address all safety issues associated with its use. The laboratory 

is responsible for maintaining a safe work environment and a current awareness file 

of OSHA regulations regarding the safe handling of the chemicals specified in this 

method. A reference file of material safety data sheets (MSDSs) shall be available to 

all personnel involved in these analyses. Additional references to laboratory safety 

can be found in Section 6.16, References 1 through 3.

6.5.4 NAF base fluids may cause skin irritation, protective gloves are recommended while 

handling these samples.

6.6 Apparatus and Materials

Note: Brand names, suppliers, and part numbers are for illustrative purposes only. No 

endorsement is implied. Equivalent performance may be achieved using apparatus and materials 

other than those specified here, but demonstration of equivalent performance meeting the 

requirements of this method is the responsibility of the laboratory.

6.6.1 Equipment for glassware cleaning.

6.6.1.1 Laboratory sink with overhead fume hood.

6.6.1.2 Kiln—Capable of reaching 450°C within 2 hours and holding 450°C 

within ±10°C, with temperature controller and safety switch (Cress 

Manufacturing Co., Santa Fe Springs, CA B31H or X31TS or equivalent).

6.6.2 Equipment for sample preparation.

6.6.2.1 Laboratory fume hood.

6.6.2.2 Analytical balance—Capable of weighing 0.1 mg.

6.6.2.3 Glassware.

a. Disposable pipettes—Pasteur, 150 mm long by 5 mm ID (Fisher 

Scientific 13–678–6A, or equivalent) baked at 400°C for a minimum 

of 1 hour.

b. Glass volumetric pipettes or gas tight syringes—1.0-mL ±1% and 0.5-

mL ±1%.

c. Volumetric flasks—Glass, class A, 10-mL, 50-mL and 100-mL.

d. Sample vials—Glass, 1- to 3-mL (baked at 400°C for a minimum of 1 

hour) with PTFE-lined screw or crimp cap.

e. Centrifuge and centrifuge tubes—Centrifuge capable of 10,000 rpm, or 

better, (International Equipment Co., IEC Centra MP4 or equivalent) 
6-4

and 50-mL centrifuge tubes (Nalgene, Ultratube, Thin Wall 25×89 

mm, #3410–2539).

6.6.3 Gas Chromatograph/Mass Spectrometer (GC/MS):

6.6.3.1 Gas Chromatograph—An analytical system complete with a temperatureprogrammable gas chromatograph suitable for split/splitless injection and 

all required accessories, including syringes, analytical columns, and gases.

a. Column—30 m (or 60 m) × 0.32 mm ID (or 0.25 mm ID) 1 μm film 

thickness (or 0.25 μm film thickness) silicone-coated fused-silica 

capillary column (J&W Scientific DB–5 or equivalent).

6.6.3.2 Mass Spectrometer—Capable of scanning from 35 to 600 amu every 1 sec 

or less, using 70 volts (nominal) electron energy in the electron impact 

ionization mode (Hewlett Packard 5970MS or comparable).

6.6.3.3 GC/MS interface—the interface is a capillary-direct interface from the GC 

to the MS.

6.6.3.4 Data system—A computer system must be interfaced to the mass 

spectrometer. The system must allow the continuous acquisition and 

storage on machine-readable media of all mass spectra obtained 

throughout the duration of the chromatographic program. The computer 

must have software that can search any GC/MS data file for ions of a 

specific mass and that can plot such ion abundance versus retention time 

or scan number. This type of plot is defined as an Extracted Ion Current 

Profile (EIP). Software must also be available that allows integrating the 

abundance in any total ion chromatogram (TIC) or EIP between specified 

retention time or scan-number limits. It is advisable that the most recent 

version of the EPA/NIST Mass Spectral Library be available.

6.7 Reagents and Standards

6.7.1 Methylene chloride—Pesticide grade or equivalent. Use when necessary for sample 

dilution.

6.7.2 Standards—Prepare from pure individual standard materials or purchase as certified 

solutions. If compound purity is 96% or greater, the weight may be used without 

correction to compute the concentration of the standard.

6.7.2.1 Crude Oil Reference—Obtain a sample of a crude oil with a known API

gravity. This oil shall be used in the calibration procedures.

6.7.2.2 Synthetic Base Fluid—Obtain a sample of clean internal olefin (IO) Lab 

drilling fluid (as sent from the supplier—has not been circulated 

downhole). This drilling fluid shall be used in the calibration procedures.

6.7.2.3 Internal standard—Prepare a 0.01 g/mL solution of 1,3,5-trichlorobenzene 

(TCB). Dissolve 1.0 g of TCB in methylene chloride and dilute to volume 
6-5

in a 100-mL volumetric flask. Stopper, vortex, and transfer the solution to 

a 150-mL bottle with PTFE-lined cap. Label appropriately, and store at 

-5°C to 20°C. Mark the level of the meniscus on the bottle to detect 

solvent loss.

6.7.2.4 GC/MS system performance test mix (SPTM) standards—The SPTM 

standards shall contain octane, decane, dodecane, tetradecane, tetradecene, 

toluene, ethylbenzene, 1,2,4-trimethylbenzene, 1-methylnaphthalene and 

1,3-dimethylnaphthalene. These compounds can be purchased individually

or obtained as a mixture (i.e., Supelco, Catalog No. 4–7300). Prepare a 

high concentration of the SPTM standard at 62.5 mg/mL in methylene 

chloride. Prepare a medium concentration SPTM standard at 1.25 mg/mL

by transferring 1.0 mL of the 62.5 mg/mL solution into a 50 mL

volumetric flask and diluting to the mark with methylene chloride. Finally, 

prepare a low concentration SPTM standard at 0.125 mg/mL by

transferring 1.0 mL of the 1.25 mg/mL solution into a 10-mL volumetric 

flask and diluting to the mark with methylene chloride.

6.7.2.5 Crude oil/drilling fluid calibration standards—Prepare a 4-point crude 

oil/drilling fluid calibration at concentrations of 0% (no spike—clean 

drilling fluid), 0.5%, 1.0%, and 2.0% by weight according to the 

procedures outlined in this section using the Reference Crude Oil:

a. Label 4 jars with the following identification: Jar 1—0%Ref-IOLab, 

Jar 2—0.5%Ref-IOLab, Jar 3—1%Ref-IOLab, and Jar 4—2%RefIOLab.

b. Weigh 4, 50-g aliquots of well mixed IO Lab drilling fluid into each of 

the 4 jars.

c. Add Reference Oil at 0.5%, 1.0%, and 2.0% by weight to jars 2, 3, and 

4 respectively. Jar 1 shall not be spiked with Reference Oil in order to 

retain a “0%” oil concentration.

d. Thoroughly mix the contents of each of the 4 jars, using clean glass 

stirring rods.

e. Transfer (weigh) a 30-g aliquot from Jar 1 to a labeled centrifuge tube. 

Centrifuge the aliquot for a minimum of 15 min at approximately

15,000 rpm, in order to obtain a solids free supernate. Weigh 0.5 g of 

the supernate directly into a tared and appropriately labeled GC 

straight vial. Spike the 0.5-g supernate with 500 μL of the 0.01g/mL

1,3,5-trichlorobenzene internal standard solution (see Section 7.2.3), 

cap with a Teflon lined crimp cap, and vortex for ca. 10 sec.

f. Repeat step 6.7.2.5(e) except use an aliquot from Jar 2.

g. Repeat step 6.7.2.5(e) except use an aliquot from Jar 3.
6-6

h. Repeat step 6.7.2.5(e) except use an aliquot from Jar 4.

i. These 4 crude/oil drilling fluid calibration standards are now used for 

qualitative and quantitative GC/MS analysis.

6.7.2.6 Precision and recovery standard (mid level crude oil/drilling fluid 

calibration standard)—Prepare a mid point crude oil/ drilling fluid 

calibration using IO Lab drilling fluid and Reference Oil at a 

concentration of 1.0% by weight. Prepare this standard according to the 

procedures outlined in Section 6.7.2.5a through 6.7.2.5e, with the 

exception that only “Jar 3” needs to be prepared. Remove and spike with 

internal standard, as many 0.5-g aliquots as needed to complete the 

GC/MS analysis (see Section 6.11.6—bracketing authentic samples every

12 hours with precision and recovery standard) and the initial 

demonstration exercise described in Section 6.9.2.

6.7.2.7 Stability of standards

a. When not used, standards shall be stored in the dark, at -5 to -20°C in 

screw-capped vials with PTFE-lined lids. Place a mark on the vial at 

the level of the solution so that solvent loss by evaporation can be 

detected. Bring the vial to room temperature prior to use.

b. Solutions used for quantitative purposes shall be analyzed within 48 

hours of preparation and on a monthly basis thereafter for signs of 

degradation. A standard shall remain acceptable if the peak area 

remains within ±15% of the area obtained in the initial analysis of the 

standard.

6.8 Sample Collection Preservation and Storage

6.8.1 Collect NAF and base fluid samples in 100- to 200-mL glass bottles with PTFE- or 

aluminum foil lined caps.

6.8.2 Samples collected in the field shall be stored refrigerated until time of preparation.

6.8.3 Sample and extract holding times for this method have not yet been established. 

However, based on initial experience with the method, samples should be analyzed 

within seven to ten days of collection and extracts should be analyzed within seven 

days of preparation.

6.8.4 After completion of GC/MS analysis, extracts shall be refrigerated at 4°C until further 

notification of sample disposal.

6.9 Quality Control

6.9.1 Each laboratory that uses this method is required to operate a formal quality 

assurance program (Section 6.16, Reference 4). The minimum requirements of this 

program shall consist of an initial demonstration of laboratory capability, and ongoing 

analysis of standards, and blanks as a test of continued performance, analyses of 
6-7

spiked samples to assess accuracy and analysis of duplicates to assess precision. 

Laboratory performance shall be compared to established performance criteria to 

determine if the results of analyses meet the performance characteristics of the 

method.

6.9.1.1 The analyst shall make an initial demonstration of the ability to generate 

acceptable accuracy and precision with this method. This ability shall be 

established as described in Section 6.9.2.

6.9.1.2 The analyst is permitted to modify this method to improve separations or 

lower the cost of measurements, provided all performance requirements 

are met. Each time a modification is made to the method, the analyst is 

required to repeat the calibration (Section 6.10.4) and to repeat the initial 

demonstration procedure described in Section 6.9.2.

6.9.1.3 Analyses of blanks are required to demonstrate freedom from 

contamination. The procedures and criteria for analysis of a blank are 

described in Section 6.9.3.

6.9.1.4 Analysis of a matrix spike sample is required to demonstrate method 

accuracy. The procedure and QC criteria for spiking are described in 

Section 6.9.4.

6.9.1.5 Analysis of a duplicate field sample is required to demonstrate method 

precision. The procedure and QC criteria for duplicates are described in 

Section 6.9.5.

6.9.1.6 Analysis of a sample of the clean NAF(s) (as sent from the supplier—has 

not been circulated downhole) used in the drilling operations is required.

6.9.1.7 The laboratory shall, on an ongoing basis, demonstrate through calibration 

verification and the analysis of the precision and recovery standard 

(Section 6.7.2.6 ) that the analysis system is in control. These procedures 

are described in Section 6.11.6.

6.9.1.8 The laboratory shall maintain records to define the quality of data that is 

generated.

6.9.2 Initial precision and accuracy—The initial precision and recovery test shall be 

performed using the precision and recovery standard (1% by weight Reference Oil in 

IO Lab drilling fluid). The laboratory shall generate acceptable precision and 

recovery by performing the following operations.

6.9.2.1 Prepare four separate aliquots of the precision and recovery standard using 

the procedure outlined in Section 6.7.2.6 . Analyze these aliquots using the 

procedures outlined in Section 6.11.
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6.9.2.2 Using the results of the set of four analyses, compute the average recovery 

(X) in weight percent and the standard deviation of the recovery(s) for 

each sample.

6.9.2.3 If s and X meet the acceptance criteria of 80% to 110%, system 

performance is acceptable and analysis of samples may begin. If, however, 

s exceeds the precision limit or X falls outside the range for accuracy, 

system performance is unacceptable. In this event, review this method, 

correct the problem, and repeat the test.

6.9.2.4 Accuracy and precision—The average percent recovery (P) and the 

standard deviation of the percent recovery (Sp) Express the accuracy

assessment as a percent recovery interval from P–2Spto P+2Sp. For 

example, if P=90% and Sp=10% for four analyses of crude oil in NAF, the 

accuracy interval is expressed as 70% to 110%. Update the accuracy

assessment on a regular basis.

6.9.3 Blanks—Rinse glassware and centrifuge tubes used in the method with 30 mL of 

methylene chloride, remove a 0.5-g aliquot of the solvent, spike it with the 500 μL of 

the internal standard solution (Section 6.7.2.3) and analyze a 1-μL aliquot of the 

blank sample using the procedure in Section 6.11. Compute results per Section 6.12.

6.9.4 Matrix spike sample—Prepare a matrix spike sample according to procedure outlined 

in Section 6.7.2.6. Analyze the sample and calculate the concentration (% oil) in the 

drilling fluid and % recovery of oil from the spiked drilling fluid using the methods 

described in Sections 6.11 and 6.12.

6.9.5 Duplicates—A duplicate field sample shall be prepared and analyzed according to 

Section 6.11. The relative percent difference (RPD) of the calculated concentrations 

shall be less than 15%.

6.9.5.1 Analyze each of the duplicates per the procedure in Section 6.11 and 

compute the results per Section 6.12.

6.9.5.2 Calculate the relative percent difference (RPD) between the two results 

per the following equation:

[6-1]

Where:

D1 = Concentration of crude oil in the sample; and

D2 = Concentration of crude oil in the duplicate sample.

6.9.5.3 If the RPD criteria are not met, the analytical system shall be judged to be 

out of control, and the problem must be immediately identified and 

corrected, and the sample batch re-analyzed.

RPD = │D1 – D2 │× 100 [(D1 + D2) /2]
6-9

6.9.6 A clean NAF sample shall be prepared and analyzed according to Section 6.11. 

Ultimately the oil-equivalent concentration from the TIC or EIP signal measured in 

the clean NAF sample shall be subtracted from the corresponding authentic field 

samples in order to calculate the true contaminant concentration (% oil) in the field 

samples (see Section 6.12).

6.9.7 The specifications contained in this method can be met if the apparatus used is 

calibrated properly, and maintained in a calibrated state. The standards used for initial 

precision and recovery (Section 6.9.2 ) and ongoing precision and recovery (Section 

6.11.6) shall be identical, so that the most precise results will be obtained. The 

GC/MS instrument will provide the most reproducible results if dedicated to the 

setting and conditions required for the analyses given in this method.

6.9.8 Depending on specific program requirements, field replicates and field spikes of 

crude oil into samples may be required when this method is used to assess the 

precision and accuracy of the sampling and sample transporting techniques.

6.10 Calibration

6.10.1 Establish gas chromatographic/mass spectrometer operating conditions given in Table 

6-1. Perform the GC/MS system hardware-tune as outlined by the manufacture. The 

gas chromatograph shall be calibrated using the internal standard technique.

Note: Because each GC is slightly different, it may be necessary to adjust the 

operating conditions (carrier gas flow rate and column temperature and temperature 

program) slightly until the retention times in Table 6-2 are met.

Table 6-1. Gas Chromatograph/Mass Spectrometer (GC/MS) Operation Conditions

Parameter Setting

Injection port 280°C

Transfer line 280°C

Detector 280°C

Initial Temperature 50°C

Initial Time 5 minutes

Ramp 50 to 300°C @ 5°C per minute

Final Temperature 300°C

Final Hold 20 minutes or until all peaks have eluted

Carrier Gas Helium

Flow rate As required for standard operation

Split ratio As required to meet performance criteria (~1:100)

Mass range 35 to 600 amu.
6-10

Table 6-2. Approximate Retention Time for Compounds

Compound Approximate retention time (minutes)

Toluene 5.6

Octane, n−C8 7.2

Ethylbenzene 10.3

1,2,4-Trimethylbenzene 16.0

Decane, −C10 16.1

TCB (Internal Standard) 21.3

Dodecane, −C12 22.9

1-Methylnaphthalene 26.7

1-Tetradecene 28.4

Tetradecane, −C14 28.7

1,3-Dimethylnaphthalene 29.7

6.10.2 Internal standard calibration procedure—1,3,5-trichlorobenzene (TCB) has been 

shown to be free of interferences from diesel and crude oils and is a suitable internal 

standard.

6.10.3 The system performance test mix standards prepared in Section 6.7.2.4 shall be used 

to establish retention times and establish qualitative detection limits.

6.10.3.1 Spike a 500-mL aliquot of the 1.25 mg/mL SPTM standard with 500 μL

of the TCB internal standard solution.

6.10.3.2 Inject 1.0 μL of this spiked SPTM standard onto the GC/MS in order to 

demonstrate proper retention times. For the GC/MS used in the 

development of this method the ten compounds in the mixture had typical 

retention times shown in Table 6-2 above. Extracted ion scans for m/z 91 

and 105 showed a maximum abundance of 400,000.

6.10.3.3 Spike a 500-mL aliquot of the 0.125 mg/mL SPTM standard with 500 μL

of the TCB internal standard solution.

6.10.3.4 Inject 1.0 μL of this spiked SPTM standard onto the GC/MS to monitor 

detectable levels. For the GC/MS used in the development of this test, all 

ten compounds showed a minimum peak height of three times signal to 

noise. Extracted ion scans for m/z 91 and 105 showed a maximum 

abundance of 40,000.

6.10.4 GC/MS crude oil/drilling fluid calibration—There are two methods of quantification: 

Total Area Integration (C8–C13) and EIP Area Integration using m/z's 91 and 105. The 

Total Area Integration method should be used as the primary technique for 

quantifying crude oil in NAFs. The EIP Area Integration method should be used as a 

confirmatory technique for NAFs. However, the EIP Area Integration method shall be 

used as the primary method for quantifying oil in enhanced mineral oil (EMO) based 

drilling fluid. Inject 1.0 μL of each of the four crude oil/drilling fluid calibration 
6-11

standards prepared in Section 6.7.2.5 into the GC/MS. The internal standard should 

elute approximately 21–22 minutes after injection. For the GC/MS used in the 

development of this method, the internal standard peak was (35 to 40)% of full scale 

at an abundance of about 3.5e+07.

6.10.4.1 Total Area Integration Method—For each of the four calibration standards 

obtain the following: Using a straight baseline integration technique, 

obtain the total ion chromatogram (TIC) area from C8 to C13. Obtain the 

TIC area of the internal standard (TCB). Subtract the TCB area from the 

C8–C13area to obtain the true C8–C13 area. Using the C8–C13 and TCB 

areas, and known internal standard concentration, generate a linear 

regression calibration using the internal standard method. The r2value for 

the linear regression curve shall be greater than or equal to 0.998. Some 

synthetic fluids might have peaks that elute in the window and would 

interfere with the analysis. In this case the integration window can be 

shifted to other areas of scan where there are no interfering peaks from the

synthetic base fluid.

6.10.4.2 EIP Area Integration—For each of the four calibration standards generate 

Extracted Ion Profiles (EIPs) for m/z 91 and 105. Using straight baseline 

integration techniques, obtain the following EIP areas:

a. For m/z 91 integrate the area under the curve from approximately 9 

minutes to 21–22 minutes, just prior to but not including the internal 

standard.

b. For m/z 105 integrate the area under the curve from approximately

10.5 minutes to 26.5 minutes.

c. Obtain the internal standard area from the TCB in each of the four 

calibration standards, using m/z 180.

d. Using the EIP areas for TCB, m/z 91 and m/z105, and the known 

concentration of internal standard, generate linear regression 

calibration curves for the target ions 91 and 105 using the internal

standard method. The r2value for each of the EIP linear regression 

curves shall be greater than or equal to 0.998.

e. Some base fluids might produce a background level that would show 

up on the extracted ion profiles, but there should not be any real peaks 

(signal to noise ratio of 1:3) from the clean base fluids.

6.11 Procedure

6.11.1 Sample Preparation

6.11.1.1 Mix the authentic field sample (drilling fluid) well. Transfer (weigh) a 30-

g aliquot of the sample to a labeled centrifuge tube.
6-12

6.11.1.2 Centrifuge the aliquot for a minimum of 15 min at approximately 15,000 

rpm, in order to obtain a solids free supernate.

6.11.1.3 Weigh 0.5 g of the supernate directly into a tared and appropriately 

labeled GC straight vial.

6.11.1.4 Spike the 0.5-g supernate with 500 μL of the 0.01g/mL 1,3,5-

trichlorobenzene internal standard solution (see Section 6.7.2.3 ), cap with 

a Teflon lined crimp cap, and vortex for ca. 10 sec.

6.11.1.5 The sample is ready for GC/MS analysis.

6.11.2 Gas Chromatography

Table 6-1 summarizes the recommended operating conditions for the GC/MS. 

Retention times for the n-alkanes obtained under these conditions are given in Table 

6-2. Other columns, chromatographic conditions, or detectors may be used if initial 

precision and accuracy requirements (Section 6.9.2 ) are met. The system shall be 

calibrated according to the procedures outlined in Section 6.10, and verified every 12 

hours according to Section 6.11.6.

6.11.2.1 Samples shall be prepared (extracted) in a batch of no more than 20 

samples. The batch shall consist of 20 authentic samples, 1 blank (Section 

6.9.3), 1 matrix spike sample (Section 6.9.4), and 1 duplicate field sample 

(6.9.5), and a prepared sample of the corresponding clean NAF used in the 

drilling process.

6.11.2.2 An analytical sequence shall be analyzed on the GC/MS where the 3 

SPTM standards (Section 6.7.2.4) containing internal standard are 

analyzed first, followed by analysis of the four GC/MS crude oil/drilling 

fluid calibration standards (Section 6.7.2.5), analysis of the blank, matrix 

spike sample, the duplicate sample, the clean NAF sample, followed by

the authentic samples.

6.11.2.3 Samples requiring dilution due to excessive signal shall be diluted using 

methylene chloride.

6.11.2.4 Inject 1.0 μL of the test sample or standard into the GC, using the 

conditions in Table 6-1.

6.11.2.5 Begin data collection and the temperature program at the time of injection.

6.11.2.6 Obtain a TIC and EIP fingerprint scans of the sample (Table 6-3).

6.11.2.7 If the area of the C8 to C13 peaks exceeds the calibration range of the 

system, dilute a fresh aliquot of the test sample weighing 0.50-g and reanalyze.
6-13

6.11.2.8 Determine the C8 to C13 TIC area, the TCB internal standard area, and the 

areas for the m/z 91 and 105 EIPs. These shall be used in the calculation 

of oil concentration in the samples (see Section 6.12).

Table 6-3. Recommended Ion Mass Numbers

Selected Ion Mass Numbers Corresponding Aromatic Compounds Typical Retention Time (Minutes)

91 Methylbenzene 6.0

Ethylbenzene 10.3

1,4-Dimethylbenzene 10.9

1,3-Dimethylbenzene 10.9

1,2-Dimethylbenzene 11.9

105 1,3,5-Trimethylbenzene 15.1

1,2,4-Trimethylbenzene 16.0

1,2,3-Trimethylbenzene 17.4

156 2,6-Dimethylnaphthalene 28.9

1,2-Dimethylnaphthalene 29.4

1,3-Dimethylnaphthalene 29.7

6.11.2.9 Observe the presence of peaks in the EIPs that would confirm the presence 

of any target aromatic compounds. Using the EIP areas and EIP linear 

regression calibrations determine approximate crude oil contamination in 

the sample for each of the target ions.

6.11.3 Qualitative Identification—See Section 6.17 of this method for schematic flowchart.

6.11.3.1 Qualitative identification shall be accomplished by comparison of the TIC 

and EIP area data from an authentic sample to the TIC and EIP area data 

from the calibration standards (see Section 6.10.4). Crude oil shall be 

identified by the presence of C10 to C13 n-alkanes and corresponding target 

aromatics.

6.11.3.2 Using the calibration data, establish the identity of the C8 to C13 peaks in 

the chromatogram of the sample. Using the calibration data, establish the 

identity of any target aromatics present on the extracted ion scans.

6.11.3.3 Crude oil is not present in a detectable amount in the sample if there are 

no target aromatics seen on the extracted ion scans. The experience of the 

analyst shall weigh heavily in the determination of the presence of peaks 

at a signal-to-noise ratio of 3 or greater.

6.11.3.4 If the chromatogram shows n-alkanes from C8 to C13 and target aromatics 

to be present, contamination by crude oil or diesel shall be suspected and 

quantitative analysis shall be determined. If there are no n-alkanes present 

that are not seen on the blank, and no target aromatics are seen, the sample 

can be considered to be free of contamination.
6-14

6.11.4 Quantitative Identification

6.11.4.1 Determine the area of the peaks from C8 to C13 as outlined in the 

calibration section (Section 6.10.4.1). If the area of the peaks for the 

sample is greater than that for the clean NAF (base fluid) use the crude 

oil/drilling fluid calibration TIC linear regression curve to determine 

approximate crude oil contamination.

6.11.4.2 Using the EIPs outlined in Section 6.10.4.2, determine the presence of any 

target aromatics. Using the integration techniques outlined in Section 

6.10.4.2, obtain the EIP areas for m/z 91 and 105. Use the crude 

oil/drilling fluid calibration EIP linear regression curves to determine 

approximate crude oil contamination.

6.11.5 Complex Samples

6.11.5.1 The most common interferences in the determination of crude oil can be 

from mineral oil, diesel oil, and proprietary additives in drilling fluids.

6.11.5.2 Mineral oil can typically be identified by its lower target aromatic content, 

and narrow range of strong peaks.

6.11.5.3 Diesel oil can typically be identified by low amounts of n-alkanes from C7

to C9, and the absence of n-alkanes greater than C25.

6.11.5.4 Crude oils can usually be distinguished by the presence of high aromatics, 

increased intensities of C8 to C13 peaks, and/ or the presence of higher 

hydrocarbons of C25 and greater (which may be difficult to see in some 

synthetic fluids at low contamination levels).

a. Oil condensates from gas wells are low in molecular weight and will 

normally produce strong chromatographic peaks in the C8–C13 range. 

If a sample of the gas condensate crude oil from the formation is 

available, the oil can be distinguished from other potential sources of 

contamination by using it to prepare a calibration standard.

b. Asphaltene crude oils with API gravity < 20 may not produce 

chromatographic peaks strong enough to show contamination at levels 

of the calibration. Extracted ion peaks should be easier to see than 

increased intensities for the C8 to C13 peaks. If a sample of asphaltene 

crude from the formation is available, a calibration standard shall be 

prepared.

6.11.6 System and Laboratory Performance

6.11.6.1 At the beginning of each 8-hour shift during which analyses are 

performed, GC crude oil/drilling fluid calibration and system performance 

test mixes shall be verified. For these tests, analysis of the medium-level 

calibration standard (1-% Reference Oil in IO Lab drilling fluid, and 1.25 
6-15

mg/mL SPTM with internal standard) shall be used to verify all 

performance criteria. Adjustments and/or re-calibration (per Section 6.10) 

shall be performed until all performance criteria are met. Only after all 

performance criteria are met may samples and blanks be analyzed.

6.11.6.2 Inject 1.0 μL of the medium-level GC/MS crude oil/drilling fluid 

calibration standard into the GC instrument according to the procedures in 

Section 6.11.2. Verify that the linear regression curves for both TIC area 

and EIP areas are still valid using this continuing calibration standard.

6.11.6.3 After this analysis is complete, inject 1.0 μL of the 1.25 mg/mL SPTM

(containing internal standard) into the GC instrument and verify the proper 

retention times are met (see Table 6-2).

6.11.6.4 Retention times—Retention time of the internal standard. The absolute 

retention time of the TCB internal standard shall be within the range 21.0 

±0.5 minutes. Relative retention times of the n-alkanes: The retention 

times of the n-alkanes relative to the TCB internal standard shall be 

similar to those given in Table 6-2.

6.12 Calculations

The concentration of oil in NAFs drilling fluids shall be computed relative to peak areas between 

C8 and C13 (using the Total Area Integration method) or total peak areas from extracted ion 

profiles (using the Extracted Ion Profile Method). In either case, there is a measurable amount of 

peak area, even in clean drilling fluid samples, due to spurious peaks and electrometer “noise” 

that contributes to the total signal measured using either of the quantification methods. In this 

procedure, a correction for this signal is applied, using the blank or clean sample correction 

technique described in American Society for Testing Materials (ASTM) Method D–3328–90, 

Comparison of Waterborne Oil by Gas Chromatography. In this method, the “oil equivalents” 

measured in a blank sample by total area gas chromatography are subtracted from that 

determined for a field sample to arrive at the most accurate measure of oil residue in the 

authentic sample.

6.12.1 Total Area Integration Method

6.12.1.1 Using C8 to C13 TIC area, the TCB area in the clean NAF sample and the 

TIC linear regression curve, compute the oil equivalent concentration of 

the C8 to C13 retention time range in the clean NAF.

Note: The actual TIC area of the C8 to C13 is equal to the C8 to C13 area 

minus the area of the TCB.

6.12.1.2 Using the corresponding information for the authentic sample, compute 

the oil equivalent concentration of the C8 to C13 retention time range in the 

authentic sample.
6-16

6.12.1.3 Calculate the concentration (% oil) of oil in the sample by subtracting the 

oil equivalent concentration (% oil) found in the clean NAF from the oil 

equivalent concentration (% oil) found in the authentic sample.

6.12.2 EIP Area Integration Method

6.12.2.1 Using either m/z 91 or 105 EIP areas, the TCB area in the clean NAF 

sample, and the appropriate EIP linear regression curve, compute the oil 

equivalent concentration of the in the clean NAF.

6.12.2.2 Using the corresponding information for the authentic sample, compute its 

oil equivalent concentration.

6.12.2.3 Calculate the concentration (% oil) of oil in the sample by subtracting the

oil equivalent concentration (% oil) found in the clean NAF from the oil 

equivalent concentration (% oil) found in the authentic sample.

6.13 Method Performance

6.13.1 Specification in this method are adopted from EPA Method 1663, Differentiation of 

Diesel and Crude Oil by GC/FID (Section 6.16, Reference 4).

6.13.2 Single laboratory method performance using an Internal Olefin (IO) drilling fluid 

fortified at 0.5% oil using a 35 API gravity oil was:

Precision and accuracy 94 ±4%

Accuracy interval—86.3% to 102%

Relative percent difference in duplicate analysis—6.2%

6.14 Pollution Prevention

6.14.1 The solvent used in this method poses little threat to the environment when recycled 

and managed properly.

6.15 Waste Management

6.15.1 It is the laboratory's responsibility to comply with all federal, state, and local 

regulations governing waste management, particularly the hazardous waste 

identification rules and land disposal restriction, and to protect the air, water, and land 

by minimizing and controlling all releases from fume hoods and bench operations. 

Compliance with all sewage discharge permits and regulations is also required.

6.15.2 All authentic samples (drilling fluids) failing the RPE (fluorescence) test (indicated 

by the presence of fluorescence) shall be retained and classified as contaminated 

samples. Treatment and ultimate fate of these samples is not outlined in this SOP.

6.15.3 For further information on waste management, consult “The Waste Management 

Manual for Laboratory Personnel”, and “Less is Better: Laboratory Chemical 

Management for Waste Reduction”, both available from the American Chemical 
6-17

Society's Department of Government Relations and Science Policy, 1155 16th Street 

NW, Washington, DC 20036.

6.16 References

1. Carcinogens—“Working With Carcinogens.” Department of Health, Education, and 

Welfare, Public Health Service, Centers for Disease Control (available through National 

Technical Information Systems, 5285 Port Royal Road, Springfield, VA 22161, 

document no. PB–277256): August 1977.

2. “OSHA Safety and Health Standards, General Industry [29 CFR 1910], Revised.” 

Occupational Safety and Health Administration, OSHA 2206. Washington, DC: January 

1976.

3. “Handbook of Analytical Quality Control in Water and Wastewater Laboratories.” 

USEPA, EMSSL-CI, EPA–600/4–79–019. Cincinnati, OH: March 1979.

4. “Method 1663, Differentiation of Diesel and Crude Oil by GC/FID, Methods for the 

Determination of Diesel, Mineral, and Crude Oils in Offshore Oil and Gas Industry

Discharges, EPA 821–R–92–008, Office of Water Engineering and Analysis Division, 

Washington, DC: December 1992.

5. U.S. EPA. 2001. Effluent Limitations Guidelines and New Source Performance 

Standards for the Oil and Gas Extraction Point Source Category. Federal Register, 66: 

6849 (22 January 2001).

6. U.S. EPA. 2001. Effluent Limitations Guidelines and New Source Performance 

Standards for the Oil and Gas Extraction Point Source Category: Correction. Federal 

Register, 66: 30811 (8 June 2001).
6-18

6.17 Schematic Flowchart for Qualitative Identification

 

Yes No

 

 No 

 

 Yes

 

 

Figure 6-1. Schematic Flowchart for Qualitative Identification

Prepare Sample for Analyses

Section 6.11.1

GC/MS Analyses 

Obtain the TIC for the Sample

Section 6.11.2

Determine the C8 to C13 TIC Area 

for comparison to calibration 

standards 

Section 6.11.2.8

Obtain the EIP for

m/z 91, m/z 105, and m/z 156

Section 6.11.2.6

C8 to C13

peaks seen on 

TIC

Peaks present for 

Target Aromatics on 

Crude oil contamination is below EIP

detection limit. 

Report not detected

Section 6.11.3.3

Integrate peaks on the EIP for comparison 

to calibration to determine approximate 

crude oil contamination 

Section 6.11.4.2
7-1

7. REVERSE PHASE EXTRACTION (RPE) METHOD FOR DETECTION OF OIL 

CONTAMINATION IN NON-AQUEOUS DRILLING FLUIDS (NAF) (EPA METHOD 1670)

7.1 Scope and Application

7.1.1 This method is used for determination of crude or formation oil, or other petroleum 

oil contamination, in non-aqueous drilling fluids (NAFs).

7.1.2 This method is intended as a positive/negative test to determine a presence of crude 

oil in NAF prior to discharging drill cuttings from offshore production platforms.

7.1.3 This method is for use in the Environmental Protection Agency's (EPA's) survey and 

monitoring programs under the Clean Water Act, including monitoring of compliance 

with the Gulf of Mexico NPDES General Permit for monitoring of oil contamination 

in drilling fluids.

7.1.4 This method has been designed to show positive contamination for 5% of 

representative crude oils at a concentration of 0.1% in drilling fluid (vol/vol), 50% of 

representative crude oils at a concentration of 0.5%, and 95% of representative crude 

oils at a concentration of 1%.

7.1.5 Any modification of this method, beyond those expressly permitted, shall be 

considered a major modification subject to application and approval of alternate test 

procedures under 40 CFR Parts 136.4 and 136.5.

7.1.6 Each laboratory that uses this method must demonstrate the ability to generate 

acceptable results using the procedure in Section 7.9.2.

7.2 Summary of Method

7.2.1 An aliquot of drilling fluid is extracted using isopropyl alcohol.

7.2.2 The mixture is allowed to settle and then filtered to separate out residual solids.

7.2.3 An aliquot of the filtered extract is charged onto a reverse phase extraction (RPE) 

cartridge.

7.2.4 The cartridge is eluted with isopropyl alcohol.

7.2.5 Crude oil contaminates are retained on the cartridge and their presence (or absence) is 

detected based on observed fluorescence using a black light.

7.3 Definitions

7.3.1 A NAF is one in which the continuous phase is a water immiscible fluid such as an 

oleaginous material (e.g., mineral oil, enhance mineral oil, paraffinic oil, or synthetic 

material such as olefins and vegetable esters).
7-2

7.4 Interferences

7.4.1 Solvents, reagents, glassware, and other sample-processing hardware may yield 

artifacts that affect results. Specific selection of reagents and purification of solvents 

may be required.

7.4.2 All materials used in the analysis shall be demonstrated to be free from interferences 

under the conditions of analysis by running laboratory reagent blanks as described in 

Section 7.9.5.

7.5 Safety

7.5.1 The toxicity or carcinogenicity of each reagent used in this method has not been 

precisely determined; however, each chemical shall be treated as a potential health 

hazard. Exposure to these chemicals should be reduced to the lowest possible level. 

Material Safety Data Sheets (MSDSs) shall be available for all reagents.

7.5.2 Isopropyl alcohol is flammable and should be used in a well-ventilated area.

7.5.3 Unknown samples may contain high concentration of volatile toxic compounds. 

Sample containers should be opened in a hood and handled with gloves to prevent 

exposure. In addition, all sample preparation should be conducted in a well-ventilated 

area to limit the potential exposure to harmful contaminants. Drilling fluid samples 

should be handled with the same precautions used in the drilling fluid handling areas 

of the drilling rig.

7.5.4 This method does not address all safety issues associated with its use. The laboratory 

is responsible for maintaining a safe work environment and a current awareness file 

of OSHA regulations regarding the safe handling of the chemicals specified in this 

method. A reference file of material safety data sheets (MSDSs) shall be available to 

all personnel involved in these analyses. Additional information on laboratory safety 

can be found in Section 7.16, References 1 and 2.

7.6 Equipment and Supplies

Note: Brand names, suppliers, and part numbers are for illustrative purposes only. No 

endorsement is implied. Equivalent performance may be achieved using apparatus and materials 

other than those specified here, but demonstration of equivalent performance that meets the 

requirements of this method is the responsibility of the laboratory.

7.6.1 Sampling equipment.

7.6.1.1 Sample collection bottles/jars—New, pre-cleaned bottles/jars, lot-certified 

to be free of artifacts. Glass preferable, plastic acceptable, wide mouth 

approximately 1–L, with Teflon-lined screw cap.

7.6.2 Equipment for glassware cleaning.

7.6.2.1 Laboratory sink.
7-3

7.6.2.2 Oven—Capable of maintaining a temperature within ±5°C in the range of 

100–250°C.

7.6.3 Equipment for sample extraction.

7.6.3.1 Vials—Glass, 25 mL and 4 mL, with Teflon-lined screw caps, baked at 

200–250°C for 1–h minimum prior to use.

7.6.3.2 Gas-tight syringes—Glass, various sizes, 0.5 mL to 2.5 mL (if spiking of 

drilling fluids with oils is to occur).

7.6.3.3 Auto pipetters—various sizes, 0.1 mL, 0.5 mL, 1 to 5 mL delivery, and 10 

mL delivery, with appropriate size disposable pipette tips, calibrated to 

within ±0.5%.

7.6.3.4 Glass stirring rod.

7.6.3.5 Vortex mixer.

7.6.3.6 Disposable syringes—Plastic, 5 mL.

7.6.3.7 Teflon syringe filter, 25-mm, 0.45 μm pore size—Acrodisc®CR Teflon 

(or equivalent).

7.6.3.8 Reverse Phase Extraction C18 Cartridge—Waters Sep-Pak®Plus, C18

Cartridge, 360 mg of sorbent (or equivalent).

7.6.3.9 SPE vacuum manifold—Supelco Brand, 12 unit (or equivalent). Used as 

support for cartridge/syringe assembly only. Vacuum apparatus not 

required.

7.6.4 Equipment for fluorescence detection.

7.6.4.1 Black light—UV Lamp, Model UVG 11, Mineral Light Lamp, Shortwave 

254 nm, or Longwave 365 nm, 15 volts, 60 Hz, 0.16 amps (or equivalent).

7.6.4.2 Black box—cartridge viewing area. A commercially available ultraviolet 

viewing cabinet with viewing lamp, or alternatively, a cardboard box or 

equivalent, approximately 14 inch × 7.5 inch × 7.5 inch; in size and 

painted flat black inside. Lamp positioned in fitted and sealed slot in 

center on top of box. Sample cartridges sit in a tray, ca. 6 inches from 

lamp. Cardboard flaps cut on top panel and side of front panel for sample 

viewing and sample cartridge introduction, respectively.

7.6.4.3 Viewing platform for cartridges. Simple support (hand made vial tray—

black in color) for cartridges so that they do not move during the 

fluorescence testing.
7-4

7.7 Reagents and Standards

7.7.1 Isopropyl alcohol—99% purity.

7.7.2 NAF—Appropriate NAF as sent from the supplier (has not been circulated 

downhole). Use the clean NAF corresponding to the NAF being used in the current 

drilling operation.

7.7.3 Standard crude oil—NIST SRM 1582 petroleum crude oil.

7.8 Sample Collection, Preservation, and Storage

7.8.1 Collect approximately one liter of representative sample (NAF, which has been 

circulated downhole) in a glass bottle or jar. Cover with a Teflon lined cap. To allow 

for a potential need to re-analyze and/or re-process the sample, it is recommended 

that a second sample aliquot be collected.

7.8.2 Label the sample appropriately.

7.8.3 All samples must be refrigerated at 0–4°C from the time of collection until extraction 

(40 CFR Part 136, Table II).

7.8.4 All samples must be analyzed within 28 days of the date and time of collection (40 

CFR Part 136, Table II).

7.9 Quality Control

7.9.1 Each laboratory that uses this method is required to operate a formal quality

assurance program (Section 7.16, Reference 3). The minimum requirements of this 

program consist of an initial demonstration of laboratory capability, and ongoing 

analyses of blanks and spiked duplicates to assess accuracy and precision and to 

demonstrate continued performance. Each field sample is analyzed in duplicate to 

demonstrate representativeness.

7.9.1.1 The analyst shall make an initial demonstration of the ability to generate 

acceptable accuracy and precision with this method. This ability is 

established as described in Section 7.9.2.

7.9.1.2 Preparation and analysis of a set of spiked duplicate samples to document 

accuracy and precision. The procedure for the preparation and analysis of 

these samples is described in Section 7.9.4.

7.9.1.3 Analyses of laboratory reagent blanks are required to demonstrate freedom 

from contamination. The procedure and criteria for preparation and 

analysis of a reagent blank are described in Section 7.9.5.

7.9.1.4 The laboratory shall maintain records to define the quality of the data that 

is generated.
7-5

7.9.1.5 Accompanying QC for the determination of oil in NAF is required per 

analytical batch. An analytical batch is a set of samples extracted at the 

same time, to a maximum of 10 samples. Each analytical batch of 10 or 

fewer samples must be accompanied by a laboratory reagent blank 

(Section 7.9.5), corresponding NAF reference blanks (Section 7.9.6), a set 

of spiked duplicate samples blank (Section 7.9.4), and duplicate analysis 

of each field sample. If greater than 10 samples are to be extracted at one 

time, the samples must be separated into analytical batches of 10 or fewer 

samples.

7.9.2 Initial demonstration of laboratory capability. To demonstrate the capability to 

perform the test, the analyst shall analyze two representative unused drilling fluids 

(e.g., internal olefin-based drilling fluid, vegetable ester-based drilling fluid), each 

prepared separately containing 0.1%, 1%, and 2% or a representative oil. Each 

drilling fluid/concentration combination shall be analyzed 10 times, and successful 

demonstration will yield the following average results for the data set:

0.1% oil—Detected in <20% of samples

1% oil—Detected in >75% of samples

2% oil—Detected in >90% of samples

7.9.3 Sample duplicates.

7.9.3.1 The laboratory shall prepare and analyze (Section 7.11.2 and 7.11.4) each 

authentic sample in duplicate, from a given sampling site or, if for

compliance monitoring, from a given discharge.

7.9.3.2 The duplicate samples must be compared versus the prepared 

corresponding NAF blank.

7.9.3.3 Prepare and analyze the duplicate samples according to procedures 

outlined in Section 7.11.

7.9.3.4 The results of the duplicate analyses are acceptable if each of the results 

give the same response (fluorescence or no fluorescence). If the results are 

different, sample non-homogenicity issues may be a concern. Prepare the 

samples again, ensuring a well-mixed sample prior to extraction. Analyze 

the samples once again.

7.9.3.5 If different results are obtained for the duplicate a second time, the 

analytical system is judged to be out of control and the problem shall be 

identified and corrected, and the samples re-analyzed.

7.9.4 Spiked duplicates—Laboratory prepared spiked duplicates are analyzed to 

demonstrate acceptable accuracy and precision.

7.9.4.1 Preparation and analysis of a set of spiked duplicate samples with each set 

of no more than 10 field samples is required to demonstrate method 

accuracy and precision and to monitor matrix interferences (interferences 
7-6

caused by the sample matrix). A field NAF sample expected to contain 

less than 0.5% crude oil (and documented to not fluoresce as part of the 

sample batch analysis) shall be spiked with 1% (by volume) of suitable 

reference crude oil and analyzed as field samples, as described in Section 

7.11. If no low-level drilling fluid is available, then the unused NAF can 

be used as the drilling fluid sample.

7.9.5 Laboratory reagent blanks—Laboratory reagent blanks are analyzed to demonstrate 

freedom from contamination.

7.9.5.1 A reagent blank is prepared by passing 4 mL of the isopropyl alcohol 

through a Teflon syringe filter and collecting the filtrate in a 4-mL glass 

vial. A Sep Pak® C18 cartridge is then preconditioned with 3 mL of 

isopropyl alcohol. A 0.5-mL aliquot of the filtered isopropyl alcohol is 

added to the syringe barrel along with 3.0 mL of isopropyl alcohol. The 

solvent is passed through the preconditioned Sep Pak®cartridge. An 

additional 2-mL of isopropyl alcohol is eluted through the cartridge. The 

cartridge is now considered the “reagent blank” cartridge and is ready for 

viewing (analysis). Check the reagent blank cartridge under the black light 

for fluorescence. If the isopropyl alcohol and filter are clean, no 

fluorescence will be observed.

7.9.5.2 If fluorescence is detected in the reagent blank cartridge, analysis of the 

samples is halted until the source of contamination is eliminated and a 

prepared reagent blank shows no fluorescence under a black light. All 

samples shall be associated with an uncontaminated method blank before 

the results may be reported for regulatory compliance purposes.

7.9.6 NAF reference blanks—NAF reference blanks are prepared from the NAFs sent from 

the supplier (NAF that has not been circulated downhole) and used as the reference 

when viewing the fluorescence of the test samples.

7.9.6.1 A NAF reference blank is prepared identically to the authentic samples. 

Place a 0.1 mL aliquot of the “clean” NAF into a 25-mL glass vial. Add 

10 mL of isopropyl alcohol to the vial. Cap the vial. Vortex the vial for 

approximately 10 sec. Allow the solids to settle for approximately 15 

minutes. Using a 5-mL syringe, draw up 4 mL of the extract and filter it 

through a PTFE syringe filter, collecting the filtrate in a 4-mL glass vial. 

Precondition a Sep Pak® C18 cartridge with 3 mL of isopropyl alcohol. 

Add a 0.5-mL aliquot of the filtered extract to the syringe barrel along 

with 3.0 mL of isopropyl alcohol. Pass the extract and solvent through the 

preconditioned Sep Pak®cartridge. Pass an additional 2-mL of isopropyl 

alcohol through the cartridge. The cartridge is now considered the NAF 

blank cartridge and is ready for viewing (analysis). This cartridge is used 

as the reference cartridge for determining the absence or presence of 

fluorescence in all authentic drilling fluid samples that originate from the 

same NAF. That is, the specific NAF reference blank cartridge is put 

under the black light along with a prepared cartridge of an authentic 

sample originating from the same NAF material. The fluorescence or 
7-7

absence of fluorescence in the authentic sample cartridge is determined 

relative to the NAF reference cartridge.

7.9.6.2 Positive control solution, equivalent to 1% crude oil contaminated mud 

extract, is prepared by dissolving 87 mg of standard crude oil into 10.00 

mL of methylene chloride. Then mix 40 μL of this solution into 10.00 mL

of IPA. Transfer 0.5 mL of this solution into a preconditioned C18

cartridge, followed by 2 ml of IPA.

7.10 Calibration and Standardization

7.10.1 Calibration and standardization methods are not employed for this procedure.

7.11 Procedure

This method is a screening-level test. Precise and accurate results can be obtained only by strict 

adherence to all details.

7.11.1 Preparation of the analytical batch.

7.11.1.1 Bring the analytical batch of samples to room temperature.

7.11.1.2 Using a large glass stirring rod, mix the authentic sample thoroughly.

7.11.1.3 Using a large glass stirring rod, mix the clean NAF (sent from the 

supplier) thoroughly.

7.11.2 Extraction.

7.11.2.1 Using an automatic positive displacement pipetter and a disposable pipette 

tip transfer 0.1-mL of the authentic sample into a 25-mL vial.

7.11.2.2 Using an automatic pipetter and a disposable pipette tip dispense a 10-mL

aliquot of solvent grade isopropyl alcohol (IPA) into the 25 mL vial.

7.11.2.3 Cap the vial and vortex the vial for ca. 10–15 seconds.

7.11.2.4 Let the sample extract stand for approximately 5 minutes, allowing the 

solids to separate.

7.11.2.5 Using a 5-mL disposable plastic syringe remove 4 mL of the extract from 

the 25-mL vial.

7.11.2.6 Filter 4 mL of extract through a Teflon syringe filter (25-mm diameter, 

0.45 μm pore size), collecting the filtrate in a labeled 4-mL vial.

7.11.2.7 Dispose of the PFTE syringe filter.

7.11.2.8 Using a black permanent marker, label a Sep Pak® C18 cartridge with the 

sample identification.
7-8

7.11.2.9 Place the labeled Sep Pak® C18 cartridge onto the head of a SPE vacuum 

manifold.

7.11.2.10 Using a 5-mL disposable plastic syringe, draw up exactly 3-mL (air free) 

of isopropyl alcohol.

7.11.2.11 Attach the syringe tip to the top of the C18 cartridge.

7.11.2.12 Condition the C18 cartridge with the 3-mL of isopropyl alcohol by

depressing the plunger slowly.

Note: Depress the plunger just to the point when no liquid remains in the 

syringe barrel. Do not force air through the cartridge. Collect the eluate in 

a waste vial.

7.11.2.13 Remove the syringe temporarily from the top of the cartridge, then remove 

the plunger, and finally reattach the syringe barrel to the top of the C18

cartridge.

7.11.2.14 Using automatic pipetters and disposable pipette tips, transfer 0.5 mL of 

the filtered extract into the syringe barrel, followed by a 3.0-mL transfer of 

isopropyl alcohol to the syringe barrel.

7.11.2.15 Insert the plunger and slowly depress it to pass only the extract and 

solvent through the preconditioned C18 cartridge.

Note: Depress the plunger just to the point when no liquid remains in the 

syringe barrel. Do not force air through the cartridge. Collect the eluate in 

a waste vial.

7.11.2.16 Remove the syringe temporarily from the top of the cartridge, then remove 

the plunger, and finally reattach the syringe barrel to the top of the C18 

cartridge.

7.11.2.17 Using an automatic pipetter and disposable pipette tip, transfer 2.0 mL of 

isopropyl alcohol to the syringe barrel.

7.11.2.18 Insert the plunger and slowly depress it to pass the solvent through the C18

cartridge.

Note: Depress the plunger just to the point when no liquid remains in the 

syringe barrel. Do not force air through the cartridge. Collect the eluate in 

a waste vial.

7.11.2.19 Remove the syringe and labeled C18 cartridge from the top of the SPE 

vacuum manifold.

7.11.2.20 Prepare a reagent blank according to the procedures outlined in Section 

7.9.5.
7-9

7.11.2.21 Prepare the necessary NAF reference blanks for each type of NAF 

encountered in the field samples according to the procedures outlined in 

Section 7.9.6.

7.11.2.22 Prepare the positive control (1% crude oil equivalent) according to Section 

7.9.6.2.

7.11.3 Reagent blank fluorescence testing.

7.11.3.1 Place the reagent blank cartridge in a black box, under a black light.

7.11.3.2 Determine the presence or absence of fluorescence for the reagent blank 

cartridge. If fluorescence is detected in the blank, analysis of the samples 

is halted until the source of contamination is eliminated and a prepared 

reagent blank shows no fluorescence under a black light. All samples must 

be associated with an uncontaminated method blank before the results may

be reported for regulatory compliance purposes.

7.11.4 Sample fluorescence testing.

7.11.4.1 Place the respective NAF reference blank (Section 7.9.6) onto the tray

inside the black box.

7.11.4.2 Place the authentic field sample cartridge (derived from the same NAF as 

the NAF reference blank) onto the tray, adjacent and to the right of the 

NAF reference blank.

7.11.4.3 Turn on the black light.

7.11.4.4 Compare the fluorescence of the sample cartridge with that of the negative 

control cartridge (NAF blank, Section 7.9.6.1) and positive control 

cartridge (1% crude oil equivalent, Section 7.9.6.2).

7.11.4.5 If the fluorescence of the sample cartridge is equal to or brighter than the 

positive control cartridge (1% crude oil equivalent, Section 7.9.6.2 ), the 

sample is considered contaminated. Otherwise, the sample is clean.

7.12 Data Analysis and Calculations

Specific data analysis techniques and calculations are not performed in this SOP.

7.13 Method Performance

This method was validated through a single laboratory study, conducted with rigorous statistical 

experimental design and interpretation (Section 7.16, Reference 4).

7.14 Pollution Prevention

7.14.1 The solvent used in this method poses little threat to the environment when recycled 

and managed properly.
7-10

7.15 Waste Management

7.15.1 It is the laboratory's responsibility to comply with all Federal, State, and local 

regulations governing waste management, particularly the hazardous waste 

identification rules and land disposal restriction, and to protect the air, water, and land 

by minimizing and controlling all releases from bench operations. Compliance with 

all sewage discharge permits and regulations is also required.

7.15.2 All authentic samples (drilling fluids) failing the fluorescence test (indicated by the 

presence of fluorescence) shall be retained and classified as contaminated samples. 

Treatment and ultimate fate of these samples is not outlined in this SOP.

7.15.3 For further information on waste management, consult “The Waste Management 

Manual for Laboratory Personnel,” and “Less is Better: Laboratory Chemical 

Management for Waste Reduction,” both available from the American Chemical 

Society's Department of Government Relations and Science Policy, 1155 16th Street, 

NW, Washington, DC 20036.

7.16 References

1. “Carcinogen—Working with Carcinogens,” Department of Health, Education, and 

Welfare, Public Health Service, Center for Disease Control, National Institute for 

Occupational Safety and Health, Publication No. 77–206, August 1977.

2. “OSHA Safety and Health Standards, General Industry,” (29 CFR 1910), Occupational 

Safety and Health Administration, OSHA 2206 (Revised, January 1976).

3. “Handbook of Analytical Quality Control in Water and Wastewater Laboratories,” 

USEPA, EMSL-Ci, Cincinnati, OH 45268, EPA–600/4–79–019, March 1979.

4. Report of the Laboratory Evaluation of Static Sheen Test Replacements—Reverse Phase 

Extraction (RPE) Method for Detecting Oil Contamination in Synthetic Based Mud 

(SBM). October 1998. Available from API, 1220 L Street, NW, Washington, DC 20005–

4070, 202–682–8000.

5. U.S. EPA. 2001. Effluent Limitations Guidelines and New Source Performance 

Standards for the Oil and Gas Extraction Point Source Category. Federal Register, 66: 

6849 (22 January 2001). 

6. U.S. EPA. 2001. Effluent Limitations Guidelines and New Source Performance 

Standards for the Oil and Gas Extraction Point Source Category: Correction. Federal 

Register, 66: 30811 (8 June 2001).
8-1

8. DETERMINATION OF THE AMOUNT OF NON-AQUEOUS DRILLING FLUID (NAF) BASE 

FLUID FROM DRILL CUTTINGS BY A RETORT CHAMBER (DERIVED FROM API

RECOMMENDED PRACTICE 13B–2) (EPA METHOD 1674)

8.1 Description

8.1.1 This procedure is specifically intended to measure the amount of non-aqueous drilling 

fluid (NAF) base fluid from cuttings generated during a drilling operation. This 

procedure is a retort test which measures all oily material (NAF base fluid) and water 

released from a cuttings sample when heated in a calibrated and properly operating 

“Retort” instrument.

8.1.2 In this retort test a known mass of cuttings is heated in the retort chamber to vaporize 

the liquids associated with the sample. The NAF base fluid and water vapors are then 

condensed, collected, and measured in a precision graduated receiver.

Note: Obtaining a representative sample requires special attention to the details of 

sample handling (e.g., location, method, frequency). See Sections 8.5 and 8.6 for 

minimum requirements for collecting representative samples. Additional sampling 

procedures in a given area may be specified by the NPDES permit controlling 

authority.

8.2 Equipment

8.2.1 Retort instrument—The recommended retort instrument has a 50-cm3volume with an 

external heating jacket.

8.2.1.1 Retort Specifications:

a. Retort assembly—retort body, cup and lid.

Material: 303 stainless steel or equivalent.

Volume: Retort cup with lid.

Cup Volume: 50- cm3.

Precision: ±0.25- cm3.

b. Condenser—capable of cooling the oil and water vapors below their 

liquification temperature.

c. Heating jacket—nominal 350 watts.

d. Temperature control—capable of limiting temperature of retort to at 

least 930°F (500°C) and enough to boil off all NAFs.

8.2.2 Liquid receiver (10- cm3, 20- cm3)—the 10- cm3and 20- cm3receivers are specially

designed cylindrical glassware with rounded bottom to facilitate cleaning and funnelshaped top to catch falling drops. For compliance monitoring under the NPDES 

program, the analyst shall use the 10- cm3liquid receiver with 0.1 ml graduations to 

achieve greater accuracy.
8-2

8.2.2.1 Receiver specifications:

Total volume: 10- cm3, 20- cm3.

Precision (0 to 100%): ±0.05 cm3, ±0.05 cm3.

Outside diameter: 10-mm, 13-mm.

Wall thickness: 1.5 ±0.1mm, 1.2 ±0.1mm.

Frequency of graduation marks (0 to 100%): 0.10- cm3, 0.10- cm3.

Calibration: To contain “TC” @ 20°C.

Scale: cm3, cm3.

8.2.2.2 Material—Pyrex®or equivalent glass.

8.2.3 Toploading balance—capable of weighing 2,000 g and precision of at least 0.1 g. 

Unless motion is a problem, the analyst shall use an electronic balance. Where motion 

is a problem, the analyst may use a triple beam balance.

8.2.4 Fine steel wool (No. 000)—for packing retort body.

8.2.5 Thread sealant lubricant: high temperature lubricant, e.g. Never-Seez®or equivalent.

8.2.6 Pipe cleaners—to clean condenser and retort stem.

8.2.7 Brush—to clean receivers.

8.2.8 Retort spatula—to clean retort cup.

8.2.9 Corkscrew—to remove spent steel wool.

8.3 Procedure

8.3.1 Clean and dry the retort assembly and condenser.

8.3.2 Pack the retort body with steel wool.

8.3.3 Apply lubricant/sealant to threads of retort cup and retort stem.

8.3.4 Weigh and record the total mass of the retort cup, lid, and retort body with steel wool. 

This is mass (A), grams.

8.3.5 Collect a representative cuttings sample (see Note in Section 8.1).

8.3.6 Partially fill the retort cup with cuttings and place the lid on the cup.

8.3.7 Screw the retort cup (with lid) onto the retort body, weigh and record the total mass. 

This is mass (B), grams.

8.3.8 Attach the condenser. Place the retort assembly into the heating jacket.

8.3.9 Weigh and record the mass of the clean and dry liquid receiver. This is mass (C), 

grams. Place the receiver below condenser outlet.
8-3

8.3.10 Turn on the retort. Allow it to run a minimum of 1 hour.

Note: If solids boil over into receiver, the test shall be rerun. Pack the retort body with 

a greater amount of steel wool and repeat the test.

8.3.11 Remove the liquid receiver. Allow it to cool. Record the volume of water recovered. 

This is (V), cm3.

Note: If an emulsion interface is present between the oil and water phases, heating the 

interface may break the emulsion. As a suggestion, remove the retort assembly from 

the heating jacket by grasping the condenser. Carefully heat the receiver along the 

emulsion band by gently touching the receiver for short intervals with the hot retort 

assembly. Avoid boiling the liquids. After the emulsion interface is broken, allow the 

liquid receiver to cool. Read the water volume at the lowest point of the meniscus.

8.3.12 Weigh and record the mass of the receiver and its liquid contents (oil plus water). 

This is mass (D), grams.

8.3.13 Turn off the retort. Remove the retort assembly and condenser from the heating jacket 

and allow them to cool. Remove the condenser.

8.3.14 Weigh and record the mass of the cooled retort assembly without the condenser. This 

is mass (E), grams.

8.3.15 Clean the retort assembly and condenser.

8.4 Calculations

8.4.1 Calculate the mass of oil (NAF base fluid) from the cuttings as follows:

8.4.1.1 Mass of the wet cuttings sample (Mw) equals the mass of the retort 

assembly with the wet cuttings sample (B) minus the mass of the empty 

retort assembly (A).

Mw = B − A [8-1]

8.4.1.2 Mass of the dry retorted cuttings (MD) equals the mass of the cooled retort 

assembly (E) minus the mass of the empty retort assembly (A).

MD = E − A [8-2]

8.4.1.3 Mass of the NAF base fluid (MBF) equals the mass of the liquid receiver 

with its contents (D) minus the sum of the mass of the dry receiver (C) and 

the mass of the water (V).

MBF = D − (C + V) [8-3]

Note: Assuming the density of water is 1 g/cm3, the volume of water is 

equivalent to the mass of the water.
8-4

8.4.2 Mass balance requirement:

The sum of MD, MBF, and V shall be within 5% of the mass of the wet sample.

[8-4]

The procedure shall be repeated if this requirement is not met.

8.4.3 Reporting oil from cuttings:

8.4.3.1 Assume that all oil recovered is NAF base fluid.

8.4.3.2 The mass percent NAF base fluid retained on the cuttings (%BFi) for the 

sampled discharge “i” is equal to 100 times the mass of the NAF base 

fluid (MBF) divided by the mass of the wet cuttings sample (Mw).

[8-5]

Operators discharging small volume NAF-cuttings discharges which do 

not occur during a NAF-cuttings discharge sampling interval (i.e., 

displaced interfaces, accumulated solids in sand traps, pit clean-out solids, 

or centrifuge discharges while cutting mud weight) shall either: (a) 

Measure the mass percent NAF base fluid retained on the cuttings 

(%BFSVD) for each small volume NAF-cuttings discharges; or (b) use a 

default value of 25% NAF base fluid retained on the cuttings.

8.4.3.3 The mass percent NAF base fluid retained on the cuttings is determined 

for all cuttings wastestreams and includes fines discharges and any 

accumulated solids discharged. (See Section 8.4.3.6 for procedures on 

measuring or estimating the mass percent NAF base fluid retained on the 

cuttings (%BF) for dual gradient drilling seafloor discharges performed to 

ensure proper operation of subsea pumps.)

8.4.3.4 A mass NAF-cuttings discharge fraction (X, unitless) is calculated for all 

NAF-cuttings, fines, or accumulated solids discharges every time a set of 

retorts is performed (Section 8.4.3.6 for procedures on measuring or 

estimating the mass NAF-cuttings discharge fraction (X) for dual gradient 

drilling seafloor discharges performed to ensure proper operation of 

subsea pumps). The mass NAF-cuttings discharge fraction (X) combines 

the mass of NAF-cuttings, fines, or accumulated solids discharged from a 

particular discharge over a set period of time with the total mass of NAFcuttings, fines, or accumulated solids discharged into the ocean during the 

same period of time (see Sections 8.5 and 8.6). The mass NAF-cuttings 

discharge fraction (X) for each discharge is calculated by direct 

measurement as:

%BFi =

MBF × 100 Mw

MD + MBF + V

= 0.95 to 1.05 Mw
8-5

[8-6]

Where:

Xi = Mass NAF-cuttings discharge fraction for NAF-cuttings, fines, or 

accumulated solids discharge “i”, (unitless)

Fi = Mass of NAF-cuttings discharged from NAF-cuttings, fines, or 

accumulated solids discharge “i” over a specified period of time 

(see Sections 8.5 and 8.6), (kg)

G = Mass of all NAF-cuttings discharges into the ocean during the 

same period of time as used to calculate Fi, (kg)

If an operator has more than one point of NAF-cuttings discharge, the 

mass faction (Xi) must be determined by: (a) Direct measurement (see

Equation 8-6); (b) using the following default values of 0.85 and 0.15 for 

the cuttings dryer (e.g., horizontal centrifuge, vertical centrifuge, squeeze 

press, High-G linear shakers) and fines removal unit (e.g., decanting 

centrifuges, mud cleaners), respectively, when the operator is only 

discharging from the cuttings dryer and the fines removal unit; or (c) using

direct measurement of “Fi” (see Equation 8-6) for fines and accumulated 

solids, using Equation 8-6A to calculate “GEST” for use as “G” in Equation 

8-6, and calculating the mass (kg) of NAF-cuttings discharged from the 

cuttings dryer (Fi) as the difference between the mass of “GEST” calculated 

in Equation 8-6A (kg) and the sum of all fines and accumulated solids 

mass directly measured (kg) (see Equation 8-6).

GEST = Estimated mass of all NAF-cuttings discharges into the ocean 

during the same period of time as used to calculate Fi

(see Equation 8-6), (kg) [8-6A]

Where:

GEST = Hole Volume (bbl) × (396.9 kg/bbl) × (1 + Z/100)

Z = The base fluid retained on cuttings limitation or standard (%) 

which apply to the NAF being discharge (see 40 CFR§§435.13. 

and 435.15).

Hole Volume 

(bbl)

= [Cross-Section Area of NAF interval (in2)] × Average 

Rate of Penetration (feet/hr) × period of time (min) used 

to calculate Fi (see Equation 8-6) × (1 hr/60 min) × 

(1 bbl/5.61 ft3) × (1 ft/12 in)2

Cross-Section 

Area of NAF 

interval (in2)

= (3.14 × [Bit Diameter (in)]2)/4

Bit Diameter 

(in)

= Diameter of drilling bit for the NAF interval producing 

drilling cuttings during the same period of time as used to 

calculate Fi (see Equation 8-6)

Average Rate 

of Penetration 

(feet/hr)

= Arithmetic average of rate of penetration into the 

formation during the same period of time as used to 

calculate Fi (see Equation 8-6)

Xi = 

Fi

G
8-6

Note: Operators with one NAF-cuttings discharge may set the mass NAFcuttings discharge fraction (Xi) equal to 1.0.

8.4.3.5 Each NAF-cuttings, fines, or accumulated solids discharge has an 

associated mass percent NAF base fluid retained on cuttings value (%BF) 

and mass NAF-cuttings discharge fraction (X) each time a set of retorts is 

performed. A single total mass percent NAF base fluid retained on 

cuttings value (%BFT) is calculated every time a set of retorts is 

performed. The single total mass percent NAF base fluid retained on 

cuttings value (%BFT) is calculated as:

%BFT,j= Σ(Xi) × (%BFi) [8-7]

Where:

%BFT,j = Total mass percent NAF base fluid retained on cuttings value for 

retort set “j” (unitless as percentage, %)

Xi = Mass NAF-cuttings discharge fraction for NAF-cuttings, fines, or 

accumulated solids discharge “i”, (unitless)

%BFi = Mass percent NAF base fluid retained on the cuttings for NAFcuttings, fines, or accumulated solids discharge “i” , (unitless as 

percentage, %)

Note: ΣXi= 1.

Operators with one NAF-cuttings discharge may set %BFT,j equal to 

%BFi.

8.4.3.6 Operators performing dual gradient drilling operations may require 

seafloor discharges of large cuttings (>1/4′) to ensure the proper operation 

of subsea pumps (e.g., electrical submersible pumps). Operators 

performing dual gradient drilling operations which lead to seafloor 

discharges of large cuttings for the proper operation of subsea pumps shall 

either: (a) Measure the mass percent NAF base fluid retained on cuttings 

value (%BF) and mass NAF-cuttings discharge fraction (X) for seafloor 

discharges each time a set of retorts is performed; (b) use the following set 

of default values, (%BF = 14%; X = 0.15); or (c) use a combination of (a) 

and (b) (e.g., use a default value for %BF and measure X).

Additionally, operators performing dual gradient drilling operations which 

lead to seafloor discharges of large cuttings for the proper operation of 

subsea pumps shall also perform the following tasks:

a. Use side scan sonar or shallow seismic to determine the presence of 

high density chemosynthetic communities. Chemosynthetic 

communities are assemblages of tube worms, clams, mussels, and 

bacterial mats that occur at natural hydrocarbon seeps or vents, 

generally in water depths of 500 meters or deeper. Seafloor discharges 
8-7

of large cuttings for the proper operation of subsea pumps shall not be 

permitted within 1,000 feet of a high density chemosynthetic 

community.

b. Seafloor discharges of large cuttings for the proper operation of subsea 

pumps shall be visually monitored and documented by a Remotely 

Operated Vehicle (ROV) within the tether limit (approximately 300 

feet). The visual monitoring shall be conducted prior to each time the 

discharge point is relocated (cuttings discharge hose) and conducted 

along the same direction as the discharge hose position. Near-seabed 

currents shall be obtained at the time of the visual monitoring.

c. Seafloor discharges of large cuttings for the proper operation of subsea 

pumps shall be directed within a 150 foot radius of the wellbore.

8.4.3.7 The weighted mass ratio averaged over all NAF well sections (%BFwell) is 

the compliance value that is compared with the “maximum weighted mass 

ratio averaged over all NAF well sections” BAT discharge limitations (see

the table in 40 CFR §435.13 and footnote 5 of the table in 40 CFR

§435.43) or the “maximum weighted mass ratio averaged over all NAF 

well sections” NSPS discharge limitations (see the table in 40 CFR

§435.15 and footnote 5 of the table in 40 CFR §435.45). The weighted 

mass ratio averaged over all NAF well sections (%BFwell) is calculated as 

the arithmetic average of all total mass percent NAF base fluid retained on 

cuttings values (%BFT) and is given by the following expression:

 

n

%BF

%BF

n

j 1

T, j

well





[8-8]

Where:

%BFwell = Weighted mass ratio averaged over all NAF well sections (unitless 

as percentage, %)

%BFT,j = Total mass percent NAF base fluid retained on cuttings value for 

retort set “j” (unitless as percentage, %)

n = Total number of retort sets performed over all NAF well sections 

(unitless)

Small volume NAF-cuttings discharges which do not occur during a NAFcuttings discharge sampling interval (i.e., displaced interfaces, 

accumulated solids in sand traps, pit clean-out solids, or centrifuge 

discharges while cutting mud weight) shall be mass averaged with the 

arithmetic average of all total mass percent NAF base fluid retained on 

cuttings values (see Equation 8-8). An additional sampling interval shall 

be added to the calculation of the weighted mass ratio averaged over all 

NAF well sections (%BFwell). The mass fraction of the small volume 

NAF-cuttings discharges (XSVD) will be determined by dividing the mass 
8-8

of the small volume NAF-cuttings discharges (FSVD) by the total mass of 

NAF-cuttings discharges for the well drilling operation (Gwell+ FSVD).

[8-9]

Where:

XSVD = Mass fraction of the small volume NAF-cuttings discharges 

(unitless)

FSVD = Mass of the small volume NAF-cuttings discharges (kg)

Gwell = Mass of total NAF-cuttings from the well (kg)

The mass of small volume NAF-cuttings discharges (FSVD) shall be 

determined by multiplying the density of the small volume NAF-cuttings 

discharges (ρsvd) times the volume of the small volume NAF-cuttings 

discharges (VSVD).

FSVD = ρsvd × VSVD [8-10]

Where:

FSVD = Mass of small volume NAF-cuttings discharges (kg)

ρsvd = Density of the small volume NAF-cuttings discharges (kg/bbl)

VSVD = Volume of the small volume NAF-cuttings discharges (bbl)

The density of the small volume NAF-cuttings discharges shall be 

measured. The volume of small volume discharges (VSVD) shall be either: 

(a) Be measured or (b) use default values of 10 bbl of SBF for each 

interface loss and 75 bbl of SBM for pit cleanout per well.

The total mass of NAF-cuttings discharges for the well (Gwell) shall be 

either: (a) Measured; or (b) calculated by multiplying 1.0 plus the 

arithmetic average of all total mass percent NAF base fluid retained on 

cuttings values [see Equation 8-8] times the total hole volume (Vwell) for 

all NAF well sections times a default value for the density the formation 

of 2.5 g/cm3(396.9 kg/bbl).

 

 

bbl

kg V bbl 396.9

n

%BF

G 1 well

n

j 1

T, j

well  

























































 



[8-11]

Where:

Gwell = Total mass of NAF-cuttings discharges for the well (kg)

 

n

%BF

n

j 1

 T, j



= see Equation 8-8 (unitless as a percentage)

Vwell = Total hole volume (Vwell) for all NAF well sections (bbl)

XSVD =

FSVD

Gwell+ FSVD
8-9

The total hole volume of NAF well sections (Vwell) will be calculated as:

[8-12]

For wells where small volume discharges associated with cuttings are 

made, %BFwell becomes:

 

 

SVD SVD

n

j 1

T, j

well SVD X %BF

n

%BF

%BF 1 X  

























































  



[8-13]

Note: See Sections 8.5 and 8.6 to determine the sampling frequency to 

determine the total number of retort sets required for all NAF well 

sections.

8.4.3.8 The total number of retort sets (n) is increased by 1 for each sampling 

interval (see Section 8.5.2.4) when all NAF cuttings, fines, or accumulated 

solids for that sampling interval are retained for no discharge. A zero 

discharge interval shall be at least 500 feet up to a maximum of three per 

day. This action has the effect of setting the total mass percent NAF base 

fluid retained on cuttings value (%BFT) at zero for that NAF sampling 

interval when all NAF cuttings, fines, or accumulated solids are retained 

for no discharge.

8.4.3.9 Operators that elect to use the Best Management Practices (BMPs) for 

NAF-cuttings shall use the procedures outlined in Section 8.6.

8.5 Requirements for Sampling Cuttings Discharge Streams for use with this Method 

8.5.1 Sampling Locations

8.5.1.1 Each NAF-cuttings waste stream that discharges into the ocean shall be 

sampled and analyzed as detailed earlier in Section Error! Reference 

source not found.Error! Reference source not found.. NAF-cuttings 

discharges to the ocean may include discharges from primary shakers, 

secondary shakers, cuttings dryer, fines removal unit, accumulated solids, 

and any other cuttings separation device whose NAF-cuttings waste is 

discharged to the ocean. NAF-cuttings wastestreams not directly

discharged to the ocean (e.g., NAF-cuttings generated from shake shakers 

and sent to a cuttings dryer for additional processing) do not require 

sampling and analysis.

8.5.1.2 The collected samples shall be representative of each NAF-cuttings 

discharge. Operators shall conduct sampling to avoid the serious 

consequences of error (i.e., bias or inaccuracy). Operators shall collect 

Vwell (barrels) = Σ

Bit diameter (in)2

× change in measured depth (ft) 1,029
8-10

NAF-cuttings samples near the point of origin and before the solids and 

liquid fractions of the stream have a chance to separate from one another. 

For example, operators shall collect shale shaker NAF-cuttings samples at 

the point where NAF-cuttings are coming off the shale shaker and not 

from a holding container downstream where separation of larger particles 

from the liquid can take place.

8.5.1.3 Operators shall provide a simple schematic diagram of the solids control 

system and sample locations to the NPDES permit controlling authority.

8.5.2 Type of Sample and Sampling Frequency

8.5.2.1 Each NAF-cuttings, fines, or accumulated solids discharge has an 

associated mass percent NAF base fluid retained on cuttings value (%BF) 

and mass NAF-cuttings discharge fraction (X) for each sampling interval 

(see Section 8.5.2.4). Operators shall collect a single discrete NAFcuttings sample for each NAF-cuttings waste stream discharged to the 

ocean during every sampling interval.

8.5.2.2 Operators shall use measured depth in feet from the Kelly bushing when 

samples are collected.

8.5.2.3 The NAF-cuttings samples collected for the mass fraction analysis (see

Equation 8-6) shall also be used for the retort analysis (see Equations 8-1 

through 8-5).

8.5.2.4 Operators shall collect and analyze at least one set of NAF-cuttings 

samples per day while discharging. Operators engaged in fast drilling (i.e., 

greater than 500 linear NAF feet advancement of drill bit per day) shall 

collect and analyze one set of NAF-cuttings samples per 500 linear NAF 

feet of footage drilled. Operators are not required to collect and analyze 

more than three sets of NAF-cuttings samples per day (i.e., three sampling 

intervals). Operators performing zero discharge of all NAF-cuttings (i.e., 

all NAF cuttings, fines, or accumulated solids retained for no discharge) 

shall use the following periods to count sampling intervals: (1) One 

sampling interval per day when drilling is less than 500 linear NAF feet 

advancement of drill bit per day; and (2) one sampling interval per 500 

linear NAF feet of footage drilled with a maximum of three sampling 

intervals per day.

8.5.2.5 The operator shall measure the individual masses (Fi, kg) and sum total 

mass (G, kg) (see Equation 8-6) over a representative period of time (e.g., 

<10 minutes) during steady-state conditions for each sampling interval 

(see Section 8.5.2.4). The operator shall ensure that all NAF-cuttings are 

capture for mass analysis during the same sampling time period (e.g., <10 

minutes) at approximately the same time (i.e., all individual mass samples 

collected within one hour of each other).
8-11

8.5.2.6 Operators using Best Management Practices (BMPs) to control NAFcuttings discharges shall follow the procedures in Section 8.6.

8.5.3 Sample Size and Handling

8.5.3.1 The volume of each sample depends on the volumetric flow rate (cm3/s) of 

the NAF-cuttings stream and the sampling time period (e.g., <10 minutes). 

Consequently, different solids control equipment units producing different 

NAF-cuttings waste streams at different volumetric flow rates will 

produce different size samples for the same period of time. Operators shall 

use appropriately sized sample containers for each NAF-cuttings waste 

stream to ensure no NAF-cuttings are spilled during sample collection. 

Operators shall use the same time period (e.g., <10 minutes) to collect 

NAF-cuttings samples from each NAF-cuttings waste stream. Each NAFcuttings sample size shall be at least one gallon. Operators shall clearly 

mark each container to identify each NAF-cuttings sample.

8.5.3.2 Operators shall not decant, heat, wash, or towel the NAF-cuttings to 

remove NAF base fluid before mass and retort analysis.

8.5.3.3 Operators shall first calculate the mass of each NAF-cuttings sample and 

perform the mass ratio analysis (see Equation 8-6). Operators with only

one NAF-cuttings discharge may skip this step (see Section 8.4.3.4).

8.5.3.4 Operators shall homogenize (e.g., stirring, shaking) each NAF-cuttings 

sample prior to placing a sub-sample into the retort cup. The bottom of the 

NAF-cuttings sample container shall be examined to be sure that solids are 

not sticking to it.

8.5.3.5 Operators shall then calculate the NAF base fluid retained on cuttings 

using the retort procedure (See Equations 8-1 through 8-5). Operators 

shall start the retort analyses no more than two hours after collecting the 

first individual mass sample for the sampling interval .

8.5.3.6 Operators shall not discharge any sample before successfully completing

the mass and retort analyses [i.e., mass balance requirements (see Section 

8.4.2) are satisfied]. Operators shall immediately re-run the retort analyses 

if the mass balance requirements (see Equation 8-4) are not within a 

tolerance of 5% (see Section 8.4.2, Equation 8-4).

8.5.4 Calculations

8.5.4.1 Operators shall calculate a set of mass percent NAF base fluid retained on 

cuttings values (%BF) and mass NAF-cuttings discharge fractions (X) for 

each NAF-cuttings waste stream (see Section 8.5.1.1) for each sampling 

interval (see Section 8.5.2.4) using the procedures outlined earlier in 

Section Error! Reference source not found.Error! Reference source 

not found..
8-12

8.5.4.2 Operators shall tabulate the following data for each individual NAFcuttings sample: (1) Date and time of NAF-cuttings sample collection; (2) 

time period of NAF-cuttings sample collection (see Section 8.5.3.1); (3) 

mass and volume of each NAF-cuttings sample; (4) measured depth (feet) 

at NAF-cuttings sample collection (see Section 8.5.2.2); (5) respective 

linear feet of hole drilled represented by the NAF-cuttings sample (feet); 

and (6) the drill bit diameter (inches) used to generate the NAF-cuttings 

sample cuttings.

8.5.4.3 Operators shall calculate a single total mass percent NAF base fluid 

retained on cuttings value (%BFT) for each sampling interval (see Section 

8.5.2.4) using the procedures outlined in Section 8.4.

8.5.4.4 Operators shall tabulate the following data for each total mass percent 

NAF base fluid retained on cuttings value (%BFT) for each NAF-cuttings 

sampling interval: (1) Date and starting and stopping times of NAFcuttings sample collection and retort analyses; (2) measured depth of well 

(feet) at start of NAF-cuttings sample collection (see Section 8.5.2.2); (3) 

respective linear feet of hole drilled represented by the NAF-cuttings 

sample (feet); (4) the drill bit diameter (inches) used to generate the NAFcuttings sample cuttings; and (5) annotation when zero discharge of NAFcuttings is performed.

8.5.4.5 Operators shall calculate the weighted mass ratio averaged over all NAF 

well sections (%BFwell) using the procedures outlined in Section 8.4.

8.5.4.6 Operators shall tabulate the following data for each weighted mass ratio 

averaged over all NAF well sections (%BFwell) for each NAF well: (1) 

Starting and stopping dates of NAF well sections; (2) measured depth 

(feet) of all NAF well sections; (3) total number of sampling intervals (see

Sections 8.5.2.4 and 8.5.2.6); (4) number of sampling intervals tabulated 

during any zero discharge operations; (5) total volume of zero discharged 

NAF-cuttings over entire NAF well sections; and (6) identification of 

whether BMPs were employed (see Section 8.6).

8.6 Best Management Practices (BMPs) for use with this Method 

8.6.1 Overview of BMPs

8.6.1.1 Best Management Practices (BMPs) are inherently pollution prevention 

practices. BMPs may include the universe of pollution prevention 

encompassing production modifications, operational changes, material 

substitution, materials and water conservation, and other such measures. 

BMPs include methods to prevent toxic and hazardous pollutants from 

reaching receiving waters. Because BMPs are most effective when 

organized into a comprehensive facility BMP Plan, operators shall develop 

a BMP in accordance with the requirements in this addendum.
8-13

8.6.1.2 The BMP requirements contained in this section were compiled from 

several Regional permits, an EPA guidance document (i.e., Guidance 

Document for Developing Best Management Practices (BMP)” (EPA 

833–B–93–004, U.S. EPA, 1993)), and draft industry BMPs. These 

common elements represent the appropriate mix of broad directions 

needed to complete a BMP Plan along with specific tasks common to all 

drilling operations.

8.6.1.3 Operators are not required to use BMPs if all NAF-cuttings discharges are 

monitored in accordance with Sections 8.1 through 8.4.

8.6.2 BMP Plan Purpose and Objectives

8.6.2.1 Operators shall design the BMP Plan to prevent or minimize the 

generation and the potential for the discharge of NAF from the facility to 

the waters of the United States through normal operations and ancillary

activities. The operator shall establish specific objectives for the control of

NAF by conducting the following evaluations.

8.6.2.2 The operator shall identify and document each NAF well that uses BMPs 

before starting drilling operations and the anticipated total feet to be 

drilled with NAF for that particular well.

8.6.2.3 Each facility component or system controlled through use of BMPs shall 

be examined for its NAF-waste minimization opportunities and its 

potential for causing a discharge of NAF to waters of the United States 

due to equipment failure, improper operation, natural phenomena (e.g., 

rain, snowfall).

8.6.2.4 For each NAF wastestream controlled through BMPs where experience 

indicates a reasonable potential for equipment failure (e.g., a tank 

overflow or leakage), natural condition (e.g., precipitation), or other 

circumstances to result in NAF reaching surface waters, the BMP Plan 

shall include a prediction of the total quantity of NAF which could be 

discharged from the facility as a result of each condition or circumstance.

8.6.3 BMP Plan Requirements

8.6.3.1 The BMP Plan may reflect requirements within the pollution prevention 

requirements required by the Minerals Management Service (see 30 CFR 

250.300) or other Federal or State requirements and incorporate any part 

of such plans into the BMP Plan by reference.

8.6.3.2 The operator shall certify that its BMP Plan is complete, on-site, and 

available upon request to EPA or the NPDES Permit controlling authority. 

This certification shall identify the NPDES permit number and be signed 

by an authorized representative of the operator. This certification shall be 

kept with the BMP Plan. For new or modified NPDES permits, the 

certification shall be made no later than the effective date of the new or 
8-14

modified permit. For existing NPDES permits, the certification shall be 

made within one year of permit issuance.

8.6.3.3 The BMP Plan shall:

a. Be documented in narrative form, and shall include any necessary plot 

plans, drawings or maps, and shall be developed in accordance with 

good engineering practices. At a minimum, the BMP Plan shall 

contain the planning, development and implementation, and 

evaluation/reevaluation components. Examples of these components 

are contained in “Guidance Document for Developing Best 

Management Practices (BMP)” (EPA 833–B–93–004, U.S. EPA, 

1993).

b. Include the following provisions concerning BMP Plan review.

• Be reviewed by permittee's drilling engineer and offshore 

installation manager (OIM) to ensure compliance with the BMP 

Plan purpose and objectives set forth in Section 8.6.2.

• Include a statement that the review has been completed and that 

the BMP Plan fulfills the BMP Plan purpose and objectives set 

forth in Section 8.6.2. This statement shall have dated signatures 

from the permittee's drilling engineer and offshore installation 

manager and any other individuals responsible for development 

and implementation of the BMP Plan.

8.6.3.4 Address each component or system capable of generating or causing a 

release of significant amounts of NAF and identify specific preventative or 

remedial measures to be implemented.

8.6.4 BMP Plan Documentation

8.6.4.1 The operator shall maintain a copy of the BMP Plan and related 

documentation (e.g., training certifications, summary of the monitoring 

results, records of NAF-equipment spills, repairs, and maintenance) at the 

facility and shall make the BMP Plan and related documentation available 

to EPA or the NPDES Permit controlling authority upon request.

8.6.5 BMP Plan Modification

8.6.5.1 For those NAF wastestreams controlled through BMPs, the operator shall 

amend the BMP Plan whenever there is a change in the facility or in the 

operation of the facility which materially increases the generation of those 

NAF-wastes or their release or potential release to the receiving waters.

8.6.5.2 At a minimum the BMP Plan shall be reviewed once every five years and 

amended within three months if warranted. Any such changes to the BMP 

Plan shall be consistent with the objectives and specific requirements 
8-15

listed in this addendum. All changes in the BMP Plan shall be reviewed by

the permittee's drilling engineer and offshore installation manager.

8.6.5.3 At any time, if the BMP Plan proves to be ineffective in achieving the 

general objective of preventing and minimizing the generation of NAFwastes and their release and potential release to the receiving waters 

and/or the specific requirements in this addendum, the permit and/or the 

BMP Plan shall be subject to modification to incorporate revised BMP 

requirements.

8.6.6 Specific Pollution Prevention Requirements for NAF Discharges Associated with 

Cuttings

8.6.6.1 The following specific pollution prevention activities are required in a 

BMP Plan when operators elect to control NAF discharges associated with 

cuttings by a set of BMPs.

8.6.6.2 Establishing programs for identifying, documenting, and repairing 

malfunctioning NAF equipment, tracking NAF equipment repairs, and 

training personnel to report and evaluate malfunctioning NAF equipment.

8.6.6.3 Establishing operating and maintenance procedures for each component in 

the solids control system in a manner consistent with the manufacturer's 

design criteria.

8.6.6.4 Using the most applicable spacers, flushes, pills, and displacement 

techniques in order to minimize contamination of drilling fluids when 

changing from water-based drilling fluids to NAF and vice versa.

8.6.6.5 A daily retort analysis shall be performed (in accordance with Sections 8.1

through 8.4) during the first 0.33 X feet drilled with NAF where X is the 

anticipated total feet to be drilled with NAF for that particular well. The 

retort analyses shall be documented in the well retort log. The operators 

shall use the calculation procedures detailed in Section 8.4 (see Equations 

8-1 through 8-8) to determine the arithmetic average (%BFwell) of the 

retort analyses taken during the first 0.33 X feet drilled with NAF.

a. When the arithmetic average (%BFwell) of the retort analyses taken 

during the first 0.33 X feet drilled with NAF is less than or equal to the 

base fluid retained on cuttings limitation or standard (see 40 CFR 

§§435.13 and 435.15), retort monitoring of cuttings may cease for that 

particular well. The same BMPs and drilling fluid used during the first 

0.33 X feet shall be used for all remaining NAF sections for that 

particular well.

b. When the arithmetic average (%BFwell) of the retort analyses taken 

during the first 0.33 X feet drilled with NAF is greater the base fluid 

retained on cuttings limitation or standard (see 40 CFR §§435.13 and 

435.15), retort monitoring shall continue for the following (second) 
8-16

0.33 X feet drilled with NAF where X is the anticipated total feet to be 

drilled with NAF for that particular well. The retort analyses for the 

first and second 0.33 X feet shall be documented in the well retort log.

• When the arithmetic average (%BFwell) of the retort analyses taken 

during the first 0.66 X feet (i.e., retort analyses taken from first and 

second 0.33 X feet) drilled with NAF is less than or equal to the 

base fluid retained on cuttings limitation or standard (see 40 CFR

§§435.13 and 435.15), retort monitoring of cuttings may cease for 

that particular well. The same BMPs and drilling fluid used during 

the first 0.66 X feet shall be used for all remaining NAF sections 

for that particular well.

• When the arithmetic average (%BFwell) of the retort analyses taken 

during the first 0.66 X feet (i.e., retort analyses taken from first and 

second 0.33 X feet) drilled with NAF is greater than the base fluid 

retained on cuttings limitation or standard (see 40 CFR §§435.13 

and 435.15), retort monitoring shall continue for all remaining 

NAF sections for that particular well. The retort analyses for all 

NAF sections shall be documented in the well retort log.

c. When the arithmetic average (%BFwell) of the retort analyses taken 

over all NAF sections for the entire well is greater that the base fluid 

retained on cuttings limitation or standard (see 40 CFR §§435.13 and 

435.15), the operator is in violation of the base fluid retained on 

cuttings limitation or standard and shall submit notification of these 

monitoring values in accordance with NPDES permit requirements. 

Additionally, the operator shall, as part of the BMP Plan, initiate a 

reevaluation and modification to the BMP Plan in conjunction with 

equipment vendors and/or industry specialists.

d. The operator shall include retort monitoring data and dates of retortmonitored and non-retort-monitored NAF-cuttings discharges 

managed by BMPs in their NPDES permit reports.

8.6.6.6 Establishing mud pit and equipment cleaning methods in such a way as to 

minimize the potential for building-up drill cuttings (including 

accumulated solids) in the active mud system and solids control equipment 

system. These cleaning methods shall include but are not limited to the 

following procedures.

a. Ensuring proper operation and efficiency of mud pit agitation 

equipment.

b. Using mud gun lines during mixing operations to provide agitation in 

dead spaces.
8-17

c. Pumping drilling fluids off of drill cuttings (including accumulated 

solids) for use, recycle, or disposal before using wash water to 

dislodge solids.
9-1

9. PAH CONTENT OF OIL BY HPLC/UV (EPA METHOD 1654, REVISION A)

9.1 Scope and Application

9.1.1 This method is designed to determine the polynuclear aromatic hydrocarbon (PAH) 

content of oil by high-performance liquid chromatography (HPLC) with an ultraviolet absorption (UV) detector. The PAH content is measured and reported as 

phenanthrene.

9.1.2 This method is for use in the Environmental Protection Agency’s (EPA’s) survey and 

monitoring programs under the Federal Water Pollution Control Act.

9.1.3 For oil in drilling muds, this method is designed to be used in conjunction with the 

extraction procedure in EPA Method 1662.

9.1.4 The level of PAH in Table 9-1 typifies the minimum level that can be detected in oil 

with this method.

9.1.5 Any modification of this method beyond those expressly permitted shall be 

considered as a major modification subject to application and approval of alternative 

test procedures under 40 CFR 136.4 and 136.5.

9.1.6 This method is restricted to use by or under the supervision of analysts experienced in 

the use of HPLC systems and in the interpretation of liquid chromatograms. Each 

analyst must demonstrate the ability to generate acceptable results with this method 

using the procedure described in Section 9.8.2.

9.2 Summary of Method

9.2.1 An oil sample is diluted in acetonitrile and a 20-μL aliquot is injected into the HPLC. 

The PAHs are partially separated by HPLC and detected with the UV detector.

9.2.2 Identification of PAH (qualitative analysis) is performed by comparing the response 

of the UV detector to the response during the retention-time range characteristic of 

the PAH in diesel oil. PAH is present when a response occurs during this retentiontime range.

9.2.3 Quantitative analysis is performed by calibrating the HPLC with phenanthrene using 

an external standard technique, and using the calibration factor to determine the 

concentration of PAH in the sample.

9.2.4 Quality is assured through reproducible calibration and testing of the extraction and 

HPLC systems.

9.3 Interferences

9.3.1 Solvents, reagents, glassware, and other sample processing hardware may lead to 

discrete artifacts and/or elevated baselines causing misinterpretation of 

chromatograms.
9-2

9.3.1.1 All materials used in the analysis shall be demonstrated to be free from 

interferences by running method blanks initially and with each sample 

batch (samples started through the extraction process at the same time, to a 

maximum of ten). Specific selection of reagents and purification of 

solvents by distillation in all-glass systems may be required.

9.3.1.2 Glassware and, where possible, reagents are cleaned by solvent rinse 

and/or baking at 450°C for a minimum of 1 hour.

9.3.2 When used in conjunction with Method 1662, blanks extracted in that method are 

treated as an integral part of this method.

9.3.3 Interferences co-extracted from samples may vary from source to source, depending 

on the diversity of the site being sampled.

9.4 Safety

9.4.1 The toxicity or carcinogenicity of each compound or reagent used in this method has 

not been precisely defined; however, each chemical should be treated as a potential 

health hazard. Exposure to these chemicals must be reduced to the lowest possible 

level.

9.4.2 The laboratory is responsible for maintaining a current awareness file of OSHA 

regulations regarding the safe handling of the chemicals specified in this method. A 

reference file of material safety data sheets (MSDSs) should also be made available to 

all personnel involved in the chemical analysis. Additional information on laboratory

safety can be found in References 1 through 3.

9.4.3 Methylene chloride has been classified as a known health hazard. All steps in this 

method which involve exposure to this compound shall be per-formed in an OSHAapproved fume hood.

9.5 Apparatus and Materials

NOTE: Brand names, suppliers, and part numbers are for illustrative purposes only. No 

endorsement is implied. Equivalent performance may be achieved using apparatus and materials 

other than those specified here, but demonstration of equivalent performance meeting the 

requirements of this method is the responsibility of the laboratory.

9.5.1 Equipment for glassware cleaning.

9.5.1.1 Laboratory sink with overhead fume hood.

9.5.1.2 Kiln: Capable of reaching 450°C within 2 hours and holding 450°C within 

±10°C, with temperature controller and safety switch (Cress 

Manufacturing Co, Sante Fe Springs, CA, B31H or X31TS, or equivalent).

9.5.2 Equipment for sample preparation.

9.5.2.1 Laboratory fume hood.
9-3

9.5.2.2 Analytical balance: Capable of weighing 0.1 mg.

9.5.2.3 Glassware.

a. Disposable pipettes: Pasteur, 150 mm long by 5 mm i.d. (Fisher 

Scientific 13-678-6A, or equivalent).

b. Glass pipettes: 1.0- and 10-mL, accurate to 1 % or better.

c. Volumetric flasks: Glass, 10- and 100-mL.

9.5.2.4 Sample vials: Amber glass, 2- to 5-mL with PTFE-lined screw-cap, to fit 

HPLC autosampler.

9.5.3 High-performance liquid chromatograph (HPLC): An analytical system complete 

with pumps, sample injector, column oven, and ultra-violet (UV) detector.

9.5.3.1 Pumping system: Capable of isocratic operation and producing a linear 

gradient from 50% water/50% acetonitrile to 100% acetonitrile in 10 

minutes (Waters 600E, or equivalent).

9.5.3.2 Sample injector: Capable of automated injection of up to 30 samples 

(Waters 700, or equivalent).

9.5.3.3 Column oven: Capable of operation at room ambient to 50°C (Waters 

TCM, or equivalent).

9.5.3.4 Column: Two C18 columns, 150 mm long by 4.6 mm i.d., 300 angstroms 

(Vydac 201 TP5415, or equivalent) connected in series, preceded by one 

C18 guard column, 30 mm long by 4.6 mm i.d., 300 angstroms (Vydac 201 

GCC54T, or equivalent), operated at the conditions shown in Table 9-1.

9.5.3.5 Detector: UV operated at 254 nm (Waters 490E, or equivalent).

9.5.4 Data system.

9.5.4.1 Data acquisition: The data system shall collect and record LC peak areas 

and retention times on magnetic media.

9.5.4.2 Calibration: The data system shall be used to calculate and maintain lists 

of calibration factors (response divided by concentration) and multi-point 

calibration curves. Computations of relative standard deviation 

(coefficient of variation) are used to test calibration linearity.

9.5.4.3 Data processing: The data system shall be used to search, locate, identify, 

and quantify the compounds of interest in each analysis. Displays of 

chromatograms are required to verify results.

9.5.4.4 Statistics on initial (Section 9.8.2) and ongoing (Section 9.12.6) 

performance shall be computed and maintained.
9-4

9.6 Reagents

9.6.1 Solvents.

9.6.1.1 Sample preparation: Methylene chloride, distilled in glass (Burdick and 

Jackson, or equivalent).

9.6.1.2 HPLC: Methanol, acetonitrile, and water, HPLC quality.

9.6.2 Standards: Purchased as solutions or mixtures with certification to their purity, 

concentration, and authenticity, or prepared from materials of known purity and 

composition. If compound purity is 96% or greater, the weight may be used without 

correction to compute the concentration of the standard. If PAH in oil from drilling 

mud is to be tested, the diesel oil standard used in this method should be from the oil 

used on the drilling rig from which the mud sample is taken. If this oil is not 

available, No. 2 diesel oil from a local source may be substituted.

9.6.2.1 Stock solutions: Prepare in methylene chloride or methanol and dilute In 

acetonitrile for injection into the HPLC. Observe the safety precautions in 

Section 9.4.

a. Diesel oil solutions

• Stock solution in methylene chloride (62.5 mg/mL): If QC extracts 

from Method 1662 are to be tested, use the oil that was spiked in 

that method. Weigh 6.25 g of diesel oil into a 100-mL groundglass-stoppered volumetric flask and fill to the mark with 

methylene chloride.

• Diesel oil calibration solution (1.25 mg/mL): After the oil in the 

stock solution (see bullet above) is completely dissolved, remove 

1.00 mL and place in a 50-mL volumetric flask. Dilute to the mark 

with acetonitrile. Mix thoroughly and transfer to a clean 150mL 

bottle with PTFE-lined cap.

b. Polynuclear aromatic hydrocarbons-naphthalene, phenanthrene, and 

indeno[1,2,3-cd]pyrene: Dissolve an appropriate amount of reference 

material in a suitable solvent. For example, weigh 10.0 mg of 

naphthalene in a 10-mL volumetric flask and fill to the mark with 

methanol. After the naphthalene is completely dissolved, transfer the 

solution to a 15-mL vial with PTFE-lined cap.

c. Stock solutions should be checked for signs of degradation prior to the 

preparation of calibration or performance test standards.

9.6.2.2 PAH calibration standards (CAL): Dilute and mix the stock solutions 

(Section 9.6.2.1b) in acetonitrile to produce the calibration standards 

shown in Table 9-2. The three solutions permit the response of 

phenanthrene to be measured as a function of concentration, and 
9-5

naphthalene and indeno[1,2,3-cd]pyrene permit the retention time window 

for PAH to be defined. The medium-level solution is used for calibration 

verification (Section 9.12.2).

9.6.2.3 Precision and recovery standard: The diesel oil calibration solution 

(Section 9.6.2.1a), second bullet) is used for initial precision and recovery

(IPR; Section 9.8.2) and ongoing precision and recovery (OPR, Section 

9.12.6).

9.6.2.4 Stability of solutions.

a. When not being used, standards are stored in the dark at -20 to -10°C 

in screw-capped vials with PTFE-lined lids. A mark is placed on the 

vial at the level of the solution so that solvent loss by evaporation can 

be detected. The vial is brought to room temperature prior to use. Any 

precipitate is redissolved and solvent is added if solvent loss has 

occurred.

b. Standard solutions used for quantitative purposes (Sections 9.6.2.1

through 9.6.2.3) shall be analyzed within 48 hours of preparation and 

on a monthly basis thereafter for signs of degradation. Standards will 

remain acceptable if the peak area remains within ±15% of the area 

obtained in the initial analysis of the standard.

9.7 Calibration

9.7.1 Assemble the HPLC and establish the operating conditions in Table 9-2.

9.7.2 Retention time adjustment.

9.7.2.1 Inject 20 μL- of the medium level calibration standard (Table 9-2).

9.7.2.2 Locate the three peaks in this standard.

9.7.2.3 Adjust the initial solvent mixture, the isocratic hold, the gradient, and the 

final isocratic hold until the retention times are within + 1 minute of the 

retention times given in Table 9-2.

9.7.3 Minimum level: Analyze 20 μL of the low-level calibration standard (Table 9-2) and 

verify that the HPLC instrument meets the minimum level for phenanthrene in Table 

9-1.

9.7.4 External standard calibration.

9.7.4.1 Analyze 20 μL of each calibration standard (Table 9-2) beginning with the 

lowest concentration and proceeding to the highest using to the procedure 

in Section 9.11.

9.7.4.2 Record the areas for the phenanthrene peak and the height of the 

phenanthrene peak in the high-level standard.
9-6

9.7.4.3 Compute the ratio of response to amount injected (calibration factor) at 

each concentration by dividing the area of the peak by the concentration of 

the standard injected. Calculate the mean of the three values to produce an 

average calibration factor.

9.7.4.4 Linearity: If the calibration factor is constant over the three point 

calibration range (< 15% relative standard deviation), linearity through the 

origin can be assumed; if not, the system shall be recalibrated.

9.7.5 The average calibration factor is verified on each working 8-hour shift by the 

measurement of the medium-level calibration standard (Section 9.12.5).

9.7.6 Single-point calibration for diesel oil: Inject the precision and recovery standard 

(Section 9.6.2.3) to produce a single calibration point for diesel oil.

9.7.6.1 Integrate the area from the retention time of naphthalene (including the 

leading edge of the naphthalene peak) through the end of the indeno[1,2,3-

cd]pyrene peak or until the detector signal returns to a stable baseline, 

whichever comes later, as shown in Figure 9-1.

9.7.6.2 Determine the calibration factor for diesel oil by dividing the integrated 

area (9.7.6.1) by the diesel oil concentration (Section 9.6.2.1a), second 

bullet).

9.8 Quality Assurance/Quality Control

9.8.1 Each laboratory that uses this method is required to operate a formal quality

assurance program (Reference 4). The minimum requirements of this program consist 

of an initial demonstration of laboratory capability, an ongoing analysis of standards 

and blanks as a test of continued performance, analyses of spiked samples to assess 

accuracy, and analysis of duplicates to assess precision. Laboratory performance is 

compared to established performance criteria to determine if the results of analyses 

meet the performance characteristics of the method. If the determination of PAH is to 

be made on extracts from Method 1662, the quality control samples for initial 

precision and recovery (IPR), spiked samples, duplicate samples, and ongoing 

precision and recovery (OPR) samples from Method 1662 shall be substituted for 

those in the QC tests below, and the specifications in Table 9-1 for extracts from 

Method 1662 shall be met.

9.8.1.1 The analyst shall make an initial demonstration of the ability to generate 

acceptable accuracy and precision with this method. This ability is 

established as described in Section 9.8.2.

9.8.1.2 The analyst is permitted to modify this method to improve separations or 

lower the costs of measurements, provided all performance requirements 

are met. Each time a modification is made to the method, the analyst is 

required to achieve the minimum level (Section 9.7.3) and to repeat the 

procedure in Section 9.8.2 to demonstrate method performance.
9-7

9.8.1.3 Analyses of spiked samples are required to demonstrate method accuracy

when extracts from Method 1662 are analyzed. The procedure and QC 

criteria for spiking are described in Section 9.8.3.

9.8.1.4 Analyses of duplicate samples are required to demonstrate method 

precision when extracts from Method 1662 are analyzed. The procedure 

and QC criteria for duplicates are described in 9.8.4.

9.8.1.5 Analyses of blanks are required to demonstrate freedom from 

contamination. The procedures and criteria for analysis of a blank are 

described in Section 9.8.5.

9.8.1.6 The laboratory shall, on an ongoing basis, demonstrate through calibration 

verification and analysis of the precision and recovery standard that the 

analysis system is in control. These procedures are described in Sections 

9.12.5 and 9.12.6.

9.8.1.7 The laboratory shall maintain records to define the quality of data that is 

generated. Development of accuracy statements is described in Sections 

9.8.3.2 and 9.12.6.4.

9.8.2 Initial precision and recovery (IPR): The initial precision and recovery test is 

performed using the precision and recovery standard. If extracts from Method 1662 

are to be analyzed, the extracts from the initial precision and recovery tests in that 

method shall be used; otherwise, the laboratory shall generate acceptable precision 

and recovery by performing the following operations.

9.8.2.1 Using diesel oil, prepare four separate aliquots of the precision and 

recovery standard (Section 9.6.2.3). If extracts from Method 1662 are 

analyzed, the extracts from the initial precision and recovery test in that 

method shall be used. Analyze these aliquots using the procedure in 

Section 9.11.

9.8.2.2 Using results of the set of four analyses, compute the average recovery (X) 

of PAH in mg/mL and the standard deviation of the recovery (s) in mg/mL 

for each aliquot by the external standard method (Sections 9.7.4 and 

9.14.4).

9.8.2.3 Compare s and X with the corresponding limits for initial precision and 

recovery in Table 9-1. If s and X meet the acceptance criteria, system 

performance is acceptable and analysis of oil samples may begin. If, 

however, s exceeds the precision limit or X falls outside the range for 

accuracy, system performance is unacceptable. In this event, review this 

method, correct the problem, and repeat the test.

9.8.3 Method accuracy: If extracts from Method 1662 are to be analyzed, the extract from 

the accuracy test in that method shall be used; otherwise, an accuracy test is 

unnecessary. The procedure for determining method accuracy is given in Section 8.3 
9-8

of Method 1662, and the specification for accuracy is given in Table 9-1 of this 

method.

9.8.3.1 Compare the percent recovery of PAH with the corresponding QC 

acceptance criteria in Table 9-1. If the results of the spike fail the 

acceptance criteria, and the recovery of the QC standard in the ongoing 

precision and recovery test (Section 9.12.6.3) is within the acceptance 

criteria in Table 9-1, an interference may be present. In this case, the result 

may not be reported for regulatory compliance purposes. If, however, the 

results of both the spike and the ongoing precision and recovery test fail 

the acceptance criteria, the analytical system is judged to be out of control 

and the problem shall be identified and corrected, and the sample batch 

reanalyzed.

9.8.3.2 As part of the QA program for the laboratory, method accuracy for 

samples shall be assessed and records shall be maintained. After the 

analysis of five spiked samples in which the recovery passes the test in 

Section 9.8.3, compute the average percent recovery (P) and the standard 

deviation of the percent recovery (P). Express the accuracy assessment as 

a percent recovery interval from P – 2Sp to P + 2 Sp. For example, if P = 

90% and Sp = 10% for five analyses of PAH in diesel oil, the accuracy

interval is expressed as 70 to 110%. Update the accuracy assessment on a 

regular basis (e.g., after each five to ten new accuracy measurements).

9.8.4 Duplicates: If extracts from Method 1662 are to be analyzed, the extracts from the 

duplicates test in that method shall be used. The procedure for preparing duplicates is 

given in Section 8.4 of Method 1662, and the specification for RPD is given in Table 

9-1 of this method. If extracts from Method 1662 are not to be analyzed, duplicates of 

the precision and recovery standard (Section 9.6.2.3) are analyzed, and the 

specification for RPD is given for PAH in diesel oil in Table 9-1 of this method.

9.8.4.1 Analyze each of the duplicates per the procedure in Section 9.11 and 

compute the results per Section 9.14.

9.8.4.2 Calculate the relative percent difference (RPD) between the two results 

per the following equation:

[9-1]

Where:

D1 = Concentration of diesel oil in the sample

D2 = Concentration of diesel oil in the second (duplicate) sample

9.8.4.3 The relative percent difference for duplicates shall meet the acceptance 

criteria in Table 9-1. If the criteria are not met, the analytical system is be 

judged to be out of control, and the problem must be immediately

identified and corrected and the sample set re-extracted and reanalyzed.

RPD = │D1 – D2 │× 100 [(D1 + D2) /2]
9-9

9.8.5 Blanks: If extracts from Method 1662 are to be analyzed, the extracts from blanks in 

that method shall be analyzed in addition to the blanks in this method.

9.8.5.1 Rinse the glassware used in preparation of the extracts in this method with 

acetonitrile and analyze a 20-μL aliquot of the rinsate using the procedure 

in Section 9.11 and compute the results per Section 9.14.

9.8.5.2 If PAH is detected in a blank at greater than the method detection limit 

(MDL) in Table 9-1, analysis of samples is halted until the source of 

contamination is eliminated and a blank shows no evidence of 

contamination.

9.8.6 The specifications contained in this method can be met if the apparatus used is 

calibrated properly, then maintained in a calibrated state. The standards used for 

initial precision and recovery (IPR, Section 9.8.2) and ongoing precision and recovery

(OPR, Section 9.12.6) should be identical, so that the most precise results will be 

obtained. The HPLC instrument will provide the most reproducible results if 

dedicated to the settings and conditions required for the analyses given in this 

method.

9.8.7 Depending on specific program requirements, field replicates and field spikes of 

diesel oil into samples may be required when Method 1662 and this method are used 

to assess the precision and accuracy of the sampling and sample transportation 

techniques.

9.9 Sample Collection, Preservation, and Handling

9.9.1 Oil samples are collected in 20- to 40-mL vials with PTFE- or aluminum-foil-lined 

caps and stored in the dark at -20 to -10°C.

9.9.2 If extracts from Method 1662 are to be analyzed, the laboratory should be aware that 

sample and extract holding times for this method have not yet been established. 

However, based on tests of wastewater for the analytes determined in this method, 

samples shall be extracted within 7 days of collection and extracts shall be analyzed 

within 40 days of extraction.

9.9.3 As a precaution against analyte and solvent loss or degradation, sample extracts are 

stored in glass bottles with PTFE-lined caps, in the dark, at -20 to -10°C.

9.10 Dilution of Oil and Extracts

9.10.1 Neat oil samples: If oil is received in neat form, it should be diluted to bring the 

concentration within the range of the instrument. If the oil is No. 2 diesel oil, the 

appropriate concentration will be approximately 2,000 μg/mL. Mineral oils and other 

oils containing a lesser PAH content will require less dilution.

9.10.2 Extracts from Method 1662: If extracts of samples from Method 1662 are to be 

analyzed, these extracts (from Section 10.4.2 of that method) are analyzed undiluted 

unless diesel oil is known or suspected to be present. Extracts of QC samples (IPR, 
9-10

OPR, matrix spikes, and duplicates) from Method 1662 are diluted by a factor of 10 

to bring them within the range of the HPLC.

9.10.3 Dilution of neat oil expected to be diesel oil.

9.10.3.1 Weigh 100 mg into a 10-mL volumetric flask and dilute to the mark with 

methylene chloride to produce a concentration of 10 mg/mL. Stopper and 

mix thoroughly.

9.10.3.2 Using a calibrated 1.0-mL volumetric pipette, withdraw 1.0 mL of the 

solution and place in a 10-mL volumetric flask. Then withdraw an 

additional 0.25 mL of the solution and place in the 10-mL volumetric flask 

(for a total of 1.25 mL). Fill to the mark with acetonitrile to produce a 

concentration of 1.25 mg/mL (1250 μg/mL). This solution will be near, 

but not above, the limit of the calibration range and will match the 

concentration of the QC samples from Method 1662 (assuming 100% 

recovery).

9.11 High-Performance Liquid Chromatography

9.11.1 Table 9-2 summarizes the recommended operating conditions for the HPLC. Included 

in this table and in Table 9-1 are retention times and the minimum level that can be 

achieved under these conditions. An example of the separation achieved for diesel oil 

by the multiple HPLC column system is shown in Figure 9-1. Other HPLC columns, 

chromatographic conditions, or detectors may be used if the requirements for the 

minimum level (Section 9.7.3) and initial precision and recovery (Section 9.8.2) are 

met.

9.11.2 Calibrate the system as described in Section 9.7 or verify calibration as described in 

Section 9.12.

9.11.3 Analysis of extracts.

9.11.3.1 Inject 20 μL of the sample extract, Method 1662 extract, or diluted QC 

extract into the HPLC using a high-pressure syringe or a constant-volume 

sample-injection loop. Record the volume injected to the nearest 0.1 μL.

9.11.3.2 Upon injection, begin the solvent program used in calibrating the column 

(Section 9.7.2.3). Record the signal from the time of injection until the 

detector returns to a stable baseline. Return the solvent to the initial 

conditions.

9.11.3.3 Using the retention-time data determined during calibration, integrate the 

area from the retention time of naphthalene (including the leading edge of 

the naphthalene peak) through the end of the indeno[1,2,3-cd]pyrene peak 

or until the detector signal returns to a stable baseline, whichever comes 

later.
9-11

9.11.4 If the height of the response during the period recorded (Section 9.11.3.2) exceeds the 

height of the response for phenanthrene during calibration (Section 9.7.4.2), dilute the 

extract by successive factors of 10 with acetonitrile and reanalyze until the response 

is within the calibration range.

9.12 HPLC System and Laboratory Performance

9.12.1 At the beginning of each 8-hour shift during which analyses are performed, HPLC 

calibration and system performance are verified. For these tests, analysis of the 

medium level calibration standard (Table 9-2) and of the diluted extract of the 

precision and recovery standard (Section 9.6.2.3) shall be used to verify all 

performance criteria. Adjustment and/or recalibration (per Section 9.7) shall be 

performed until all performance criteria are met. Only after all performance criteria 

are met may samples and blanks be analyzed.

9.12.2 Inject 20 μL of the medium-level calibration standard (Table 9-2) into the HPLC 

instrument according to the procedure in Section 9.11.

9.12.3 Retention time: The absolute retention times of the naphthalene, phenanthrene, and 

indeno[1,2,3-cd]pyrene peaks shall be within ±30 seconds of the respective retention 

times in the initial calibration (Section 9.7.2.3).

9.12.4 HPLC resolution: Resolution is acceptable if the peak width at half-height of the 

phenanthrene peak is less than 30 seconds.

9.12.5 Calibration verification: Compute the concentration of phenanthrene based on the 

average calibration factor (Section 9.7.4.4). The concentration shall be within the 

limits in Table 9-1. If calibration is verified, system performance is acceptable and 

analysis of blanks and QC samples may begin. If, however, the concentration falls 

outside of the calibration verification range, system performance is unacceptable. In 

this case, correct the problem and repeat the test, or recalibrate (Section 9.7.4).

9.12.6 Ongoing precision and recovery (OPR): If the extract is from Method 1662, the OPR 

standard from that method shall be used and the specification for the OPR from 

Method 1662 in Table 9-1 shall be met; if not, a sample of diesel oil shall be diluted 

per the procedure in Section 9.10 and shall be used for the OPR test.

9.12.6.1 Analyze the appropriate OPR standard.

9.12.6.2 Compute the concentration of PAH in this standard per Section 9.14.

9.12.6.3 Compare the concentration with the limits for ongoing precision and 

recovery in Table 9-1. If the concentration is in the range specified, the 

analytical processes are in control and analysis of blanks and samples may

proceed. If, however, the concentration is not in the specified range, these 

processes are not in control. In this event, correct the problem, re-extract 

the sample batch if the OPR is from Method 1662, or redilute the oil 

sample (per Section 9.10.3). and repeat the ongoing precision and 

recovery test.
9-12

9.12.6.4 Add results which pass the specification in Section 9.12.6.3 to initial and 

previous ongoing data. Update QC charts to form a graphic representation 

of continued laboratory performance. Develop a statement of laboratory

data quality for each analyte by calculating the average percent recovery

(R) and the standard deviation of percent recovery (Sr). Express the 

accuracy as a recovery interval from R – 2Sr, to R + 2 Sr. For example, if 

R = 95% and Sr = 5%, the accuracy is 85 to 105%.

9.13 Qualitative Identification

9.13.1 Qualitative determination is accomplished by comparison of data from analysis of a 

sample or blank with data from analysis of the calibration verification standard 

(Section 9.12.5).

9.13.2 PAH is identified in the sample by the presence of peaks and/or an elevated baseline 

(hump) between the retention times of the naphthalene and indeno[1,2,3-cd]pyrene 

peaks (Section 9.11.3.3), as shown in Figure 9-1. The experience of the analyst shall 

weigh heavily in interpretation of the chromatogram.

9.14 Quantitative Determination

9.14.1 Using the data system, compute the concentration of the PAH detected in the solution 

injected into the HPLC (in μg/mL) using the calibration factor (Section 9.7.4).

9.14.2 Concentration of PAH in oil: If neat oil was analyzed, the concentration of PAH in 

the oil is determined using the following equation:

[9-2]

Where:

Co = Concentration of PAH in the oil sample

Cp = Concentration of PAH measured (from Sections 9.11.4 and 9.14.1)

Ci = Concentration of oil in the solution injected into the HPLC (from

Sections 9.10.3.2, 9.11.4, and 9.14.1)

9.14.3 Concentration of diesel oil in QC extracts from Method 1662: Calculate the 

concentration of diesel oil in QC extracts from Method 1662 by integrating the area 

per Section 9.7.6.1 and using the calibration from Section 9.7.6.2 of this method, 

taking into account the dilution of these extracts (Section 9.10.2).

9.14.4 Concentration of PAH in oil from Method 1662: The PAH content of oil is 

complicated by the splitting and possible dilution of these extracts.

9.14.4.1 Concentration in undiluted extracts: This concentration is determined by 

Equation 9-3:

Co = [9-3]

Ve × Cp

=

5 × I Cp

1/5 × WT WT

Co (mg/g) = Cp (μg/mL)

Ci (mg/mL)
9-13

Where:

Co = Concentration of PAH in the oil sample

Ve = Amount of extract split for HPLC analysis, in mL (1.0 mL)

Cp = Concentration of PAH measured

WT = Weight of oil in the concentration tube in Method 1662 (Section 

11.5.5 of Method 1662)

1/5 = Fraction of this weight used for the PAH determination

9.14.4.2 Concentration in diluted extracts: If the extract was diluted by a factor of 

10 (Section 9.10.3 or 9.11.4), the concentration determined in Section 

9.14.4.1 is multiplied by 10.

9.14.5 If the concentration is to be expressed as weight percent, Co is multiplied by 0.1.

9.14.6 Report results to three significant figures without correction for recovery.

9.15 Method Performance

This method was validated in a single laboratory (Reference 6) using samples of hot-rolled 

drilling mud (Reference 7).

9.16 References

1. “Carcinogens-Working With Carcinogens.” Department of Health, Education, and 

Welfare, Public Health Service, Centers for Disease Control [available through National 

Technical Information System, 5285 Port Royal Road, Springfield, VA 22161, document 

no. PB2772561: August 1977.

2. “OSHA Safety and Health Standards, General Industry [29 CFR 1910], Revised.” 

Occupational Safety and Health Administration, OSHA 2206. Washington, DC: January 

1976.

3. “Safety in Academic Chemistry Laboratories (3rd Edition).” American Chemical Society

Publication, Committee on Chemical Safety. Washington, DC: 1979.

4. “Handbook of Analytical Quality Control in Water and Wastewater Laboratories.” 

USEPA, EMSL-Ci, EPA-600/4-79-019. Cincinnati, OH: March 1979.

5. “Standard Practice for Sampling Water,” ASTM Annual Book of Standards, Part 31, 

D337076, ASTM. Philadelphia, PA: 1980.

6. “Determination of Polynuclear Aromatic Hydrocarbons and Diesel by Modified EPA 

Method 9310.” Prepared for the American Petroleum Institute c/o Shell Development Co, 

Westhollow Research Center, 3333 Highway 6 South, Houston, TX 77082 by Analytical 

Technologies Inc., 225 Commerce Drive, Fort Collins, CO 80524: March 29 1991, 

April 12, 1991, and August 18, 1992.

7. “Results of the API Study of Extraction and Analysis Procedures for the Determination 

of Diesel Oil in Drilling Muds (Final Report).” American Petroleum Institute, Offshore 
9-14

Effluent Guidelines Steering Committee, Technology Work Group, Prepared by I.C. 

Raia, Shell Development Co. Houston, TX: April 18, 1991.

Table 9-1. Performance Data and Method Acceptance Criteria for PAH

Criterion Units PAH in Diesel Oil a

Diesel Oil in Mud 

Extract b Phenanthrene

Minimum level cμg/mL 100 — 0.1

Method Detection Limit dμg/mg 7.6 — —

Initial prec and recov

Precision (std dev)

PAH in diesel oil e mg/mL 120 — —

Diesel in mud extract f mg/mL — 0.55 —

Recovery

PAH in diesel oil e mg/mL 1,090 – 1,340 — —

Diesel in mud extract f mg/mL — 0.84 – 1.95 —

Calibration verification gμg/mL — — 0.39 – 0.61

Ongoing prec and recov

PAH in diesel oil e mg/mL 1,010 – 1,450 — —

Diesel in mud extract f mg/mL — 0.76 – 2.15 —

Matrix spike recovery fpct — 0.43 – 2.39 —

Duplicates RPD 9.5 44 —

a CAS Registry number 68534-30-5; No. 2 diesel oil used for these tests.

b

From Method 1662.

c This is a minimum level at which the analytical system shall give recognizable signals and acceptable calibration 

points..

d

40 CFR Part 136, Appendix B; MDL is measured as PAH in oil.

e Test concentration of diesel oil = 1,250 μg/mL.

f Test concentration in diluted extract = 1.25 mg/mL.

g Test concentration = 0.50 μg/mL.

Table 9-2. HPLC Calibration Data

Analyte

Retention Time a

(minutes)

Calibration Solution Concentration (μg/mL)

Low Medium High

Naphthalene 7.6 — 5 —

Phenanthrene 10.3 0.1 0.5 2.0

Indeno[l23-cd]pyrene 18.9 — 0.5 —

Diesel oil b7.4 – 20.0 100 400 2,000

a Column system: Two C18 columns (150 mm long by 4.6 mm i.d., 300 angstroms) connected in series,

preceded by one C18 guard column (30 mm long by 4.6 mm i.d., 300 angstroms). Column temperature 30°C;

solvent flow rate 1.5 mL/min; linear gradient from 50% water/50% acetonitrile at injection to 100%

acetonitrile in 10 minutes, hold at 100% acetonitrile for 15 minutes.

b Diesel oil is calibrated separately using a single point calibration (Section 9.7.6).
9-15

Figure 9-1. Liquid Chromatography of the Three-Component Standard and of No. 2 Diesel 

Oil
10-1

10. METHOD FOR CONDUCTING A SEDIMENT TOXICITY TEST WITH LEPTOCHEIRUS 

PLUMULOSUS AND NON-AQUEOUS DRILLING FLUIDS OR SYNTHETIC-BASED DRILLING 

MUDS (EPA METHOD 1644)

10.1 Summary of Method 

This test method describes the procedures for obtaining data regarding the effects of nonaqueous drilling fluids (NAFs) or synthetic based drilling muds (SBMs) on the marine 

amphipod, Leptocheirus plumulosus. EPA is regulating the sediment toxicity in NAFs 

and SBMs that are discharged from oil and gas extraction facilities in coastal and 

offshore waters as an indicator for toxic pollutants (see 40 CFR 435.13, 435.15, 435.43, 

and 435.45). EPA established the use of this test and related limits to encourage the use 

of less toxic drilling fluids and additives. The sediment toxicity of the NAF-cuttings at 

the point of discharge is measured by this modified sediment toxicity test using a natural 

sediment and Leptocheirus plumulosus as the test organism.

EPA promulgated the use of this method with revisions to the Oil and Gas Extraction 

effluent guidelines (U.S. EPA, 2001). This initial method was consistent with ASTM 

Standard Guide E 1367-92 (ASTM 1997). Subsequent to this rulemaking EPA updated 

this method in the NPDES permit for the Western Gulf of Mexico (GMG290000, 

Appendix A) to be consistent with ASTM E 1367-99 (ASTM 2000). This method is a republication of the sediment toxicity test in NPDES permit for the Western Gulf of 

Mexico (GMG290000, Appendix A).

10.2 Test Requirements and Materials

10.2.1 Test Species (Leptocheirus plumulosus)

10.2.1.1 Description: Leptocheirus plumulosus is an infaunal amphipod that is 

indigenous to subtidal regions along the east coast of the U.S. This 

amphipod constructs U-shaped burrows in the top 5 cm of fine sand to 

silty clay sediments (ASTM E 1367-99). As a result of its broad salinity

and particle size tolerances, it is a desirable test species for a variety of 

toxicity testing programs.

10.2.1.2 Collection and Handling: In the field, amphipods can be collected using 

sediment grab samplers such as Peterson and Ponar dredges. This species 

has been collected in various tributaries of the Chesapeake Bay for various 

toxicity testing programs (ASTM E 1367-99). The contents of each grab 

should be sieved through a 500 mesh screen. The sediment and organisms 

retained on the screen are gently rinsed into plastic buckets containing 

sediment and water from the collection site. These buckets are quickly 

transported back to the laboratory and aerated. See ASTM E 1367-99 for 

more details on collection and handling.

10.2.1.3 Holding and Acclimation: Amphipods can be placed in aquaria containing 

a 1-2 cm deep layer of collection site sediment that has been sieved 

through a 500 mesh screen. Amphipod density should be about 200-300 

per 40 L aquarium with vigorous aeration. Two to three days are sufficient 
10-2

for acclimation to test conditions, and during this period a gradual change 

over from site water to test water is recommended (ASTM E 1367-99).

10.2.1.4 Environmental Tolerances: L. plumulosus is tolerant of a broad salinity 

range, from near 0 to 33 g/kg (‰) (ASTM E 1367- 99). This species has 

demonstrated up to 100% survival in >90% silt-clay sediment and an 

average of 85% survival in >95% sand/gravel sediment (ASTM E 1367-

99). The ASTM data are consistent with data published from other studies 

indicating that L. plumulosus is tolerant of sandy and silty sediments. For 

example, Schlekat et al. (1992) noted a mean survival of 97.5% when L. 

plumulosus was exposed for 10 days to field collected sediments ranging 

from 98.1% sand to 96.5% fines. Further, this species was collected in the 

field in sediments consisting of 99.9% sand and 92.1% fines, indicating 

that L. plumulosus is a generalist and can thrive in a variety of sediment 

types (Schlekat et al. 1992).

However, the fine fraction of sediments in the Schlekat et al. study did not 

exceed 55% clay, indicating that the fine fraction was a mixture of silt and 

clay sized particles. Data from other studies indicated that this species is 

intolerant of sediments high in clay content. McGee et al. (1999) noted 

acceptable survival when this species was exposed to Baltimore Harbor 

sediments containing up to 72% clay. However, Emery et al. (1997) noted 

significantly reduced amphipod survival when L. plumulosus was exposed 

for 10 days to Magothy River, Maryland sediment (amended with beach 

sand and kaolinite clay) containing 84%, 90%, and 100% clay.

These data indicated that the tolerance range of this amphipod to clay

content is between about 72 to 84%. As such, caution should be used 

when conducting L. plumulosus toxicity tests with sediments with clay

content greater than about 70%. This should not have a significant impact 

on using this species in the NAF and SBM toxicity testing program, since 

field sediments seldom exceed 70% clay content (Suedel and Rodgers, 

1991).

10.2.2 Culture Methods (Leptocheirus plumulosus) 

10.2.2.1 Overview: Populations of L. plumulosus can be maintained through 

several generations in the laboratory. The culture conditions specified in 

ASTM E 1367-92 and E 1367-99 are provided in Table 1. 
10-3

Table 10-1. Culture conditions for L. plumulosus. Conditions listed are consistent with 

culture conditions specified in ASTM E 1367-92 and subsequent updates (E 1367-99)

Parameter Conditions

Temperature 20±1 oC 

Salinity 20±1‰

Light Quality Wide-spectrum fluorescent or cool white lights

Illuminance 500-1,000 lux

Photoperiod 14h light: 10h dark

Culture Chamber Shallow plastic tubs or glass aquaria

Sediment Volume 1-2 cm depth at bottom of each culture chamber

Renewal of overlying water Static renewal (30-50% water volume change 2-4

times per week)

Number of 

organisms/chamber

Start with about 300 mixed age (mostly immature and 

young adults) individuals per chamber

Feeding 0.1 to 0.5 g dry mixture 2-3 times per week (see section 

10.2.2.2)

Aeration Continuous gentle to moderate aeration so as to not

suspend sediments

Overlying Water Clean natural or synthetic seawater

Overlying water quality Salinity, temperature, and ammonia during culture

start-up

In addition to the conditions provided in Table 1, there are other 

conditions that are important in maintaining healthy L. plumulosus

cultures, including identifying a source of clean sediment, sieving 

sediments before use, and the quality of the raw materials used to prepare 

their food. Preferably, the sediment and water used to culture the 

amphipods should be collected from the same area as those used in NAF 

tests. Fine-grained sediments have been shown to be suitable for this 

purpose (ASTM E 1367-92). Sediments collected in the field for culturing 

purposes should be first sieved through a 2,000 m mesh sieve to remove 

large debris and then through a 500 m mesh sieve to remove any 

indigenous organisms. L. plumulosus cultures should be maintained at 

20±1 oC and 20±1‰ salinity. If used, natural seawater should be filtered 

through a 5 micron filter before adding to cultures. New culture chambers 

should be aerated and allowed to equilibrate overnight before adding 

amphipods. Water used to start a new culture chamber should be renewed 

24 h after initiation and before amphipods are added to culture chambers; 

otherwise, culture water should be renewed in conjunction with feeding.

Cultures should be observed daily to ensure sufficient aeration. An 

abundance of amphipods on the sediment surface during daylight hours 

may indicate insufficient dissolved oxygen or overcrowding, as 

amphipods typically remain in their burrows unless they are searching for 

food or a mate. Culture chambers should be terminated and restarted with 

fresh sediment about once every 8 weeks to avoid overcrowding. 
10-4

Overcrowding may lead to stress due to food or space limitations, and may

also result in reduced female fecundity, thus reducing the relative health of 

the population of amphipods in a given culture chamber.

Cultures should be routinely inspected for the presence of indigenous 

worms and copepods, a microbial build-up, or black and sulfurous 

conditions beneath the sediment surface. Microbial growth appears as a 

white or gray growth associated with uneaten food, and is indicative of 

overfeeding. Presence of indigenous species, excess microbial growth, or 

black and sulfurous conditions may necessitate discarding the affected 

culture chamber.

10.2.2.2 Feeding: A mixture of micro-algae, yeast, fish food flakes, alfalfa powder, 

ground cereal leaves, and shrimp maturation feed has been used to feed 

cultures (ASTM E 1367-92 and E 1367-99). Micro-algae used in culturing 

include Pseudoisochrysis paradoxa, Phaeodactylum tricornutum, and 

Tetraselmis suecica mixed in equal parts on a volume basis. These algae 

provide a source of fatty acids that may otherwise be absent in the diet. In 

practice, however, it should be noted that L. plumulosus has been cultured 

successfully without the algal mixture and the yeast. The dry food portion 

of the diet that has been used to successfully culture L. plumulosus is 

shown in Table 2. 

Table 10-2. Dry food portion of the diet that has been used to successfully culture L. 

plumulosus

Dietary Component Proportion

Fish food flakes (e.g., TetraMin®) 48.0%

Alfalfa powder 24%

Ground cereal leaves (dried wheat leaves) 24%

Shrimp maturation feed (e.g., Neo-Novum®) 4.0%

This dry food mixture should be homogenized into a fine powder and fed 

to each culture chamber at a rate of 0.1 to 0.5 g two to three times per 

week, depending on culture densities. Overfeeding may result in microbial 

build-up on the sediment surface. The quality of the alfalfa powder and 

dried wheat leaves may not be consistent among suppliers, thus potentially

adversely affecting culture performance. Feeding should occur 

immediately after culture water changes.

10.2.3 Obtaining Amphipods for Starting a Test (Leptocheirus plumulosus)

10.2.3.1 Immature and adult amphipods of mixed sexes and approximately 3 to 5 

mm in length (as measured from the base of the first antenna to the end of 

the third pleon segment along the dorsal surface) are used in toxicity tests, 

as they are easier to handle and count than younger individuals. Gravid 

females are not used in testing. 
10-5

10.2.3.2 The 3 to 5 mm size class individuals are passed through a 1,000 m mesh 

sieve and are retained on a 710 m mesh sieve. A 500 m mesh sieve has 

been used previously to retain amphipods of the size needed, but this will 

result in a wider size range of amphipods used for testing. 

10.2.3.3 In preliminary NAF experiments, this wide size range may have 

contributed to variability in mortality observed that was not present when 

the 710 m mesh sieve was used to retain amphipods in later experiments. 

The amphipods passing through a 1000 m mesh sieve but trapped on a 710 

m mesh sieve provide a more uniform size range of animals that is thought 

to decrease the previously-observed variability in mortality. Laboratories 

are encouraged to use this type of approach to reduce the variability in the 

size of amphipods used in the NAF/SBM testing program.

10.2.4 Control Sediments

10.2.4.1 Overview: Control sediment must meet certain minimum requirements to 

be used in the NAF/SBM testing program. The primary requirement is that 

the sediment should be able to support L. plumulosus in cultures for 

extended periods of time. This will ensure that the sediment is chemically

nontoxic and that the physical and chemical characteristics of the sediment 

(e.g., total organic carbon, particle size distribution, and moisture content) 

are within the tolerance range of the test species. It is expected that 

separate aliquots of the culture sediment will also be used as a control 

sediment to be amended by NAFs or SBMs in the NAF/SBM testing 

program. Any modifications made to the control sediments should be 

noted in the report.

10.2.4.2 Characterization: Sediments used in testing should be characterized for 

total organic carbon (TOC), particle size distribution (sand, silt, and clay), 

and percent water content. These parameters have been shown to influence 

the results of NAF/SBM toxicity to L. plumulosus in initial experiments. 

Variations in these sediment characteristics should be quantified so that 

potential effects of these parameters on test results can be closely

monitored. 

10.2.4.3 Collection: Control sediments should be collected from the amphipod 

collection site or from another area that can provide a consistent source of 

sediment with characteristics within the tolerance range of L. plumulosus. 

Sediments showing evidence of chemical contamination should not be 

used in the NAF/SBM testing program. Any site water overlying the 

sediment should be retained so that fine particles suspended in the water 

can be re-combined with the sediment before use. Sediment salinity and 

temperature should be recorded at the time of collection. Sediment 

collected for use should be homogenized and a composite sample prepared 

for analysis for the parameters outlined above.

10.2.4.4 Sieving: Sediments collected in the field for culturing and testing purposes 

should be first press-sieved through a 2,000 or similar mesh sieve to 
10-6

remove large debris and then through a 500 mesh sieve to remove any 

indigenous organisms. Sediments have also been press-sieved through a 

250 to 350 mesh sieve prior to testing to aid in the enumeration of 

amphipods on a 500 mesh sieve at test termination.

10.2.4.5 Storage: The control sediment should be stored in plastic or glass 

containers at 4 oC until test initiation. The sediment should be stored in the 

dark and should not be allowed to freeze or dry out during storage (ASTM 

E 1367-92).

10.2.5 Test Water

10.2.5.1 Water used in the NAF/SBM testing program should be available in 

sufficient quantities and be acceptable to L. plumulosus. The minimum 

requirement for acceptable water for use in the NAF/SBM testing program

is that healthy test organisms survive in the water, and in the water plus 

control sediment, for the duration of holding and testing without showing 

signs of disease or stress (ASTM E 1367-99). Another test for 

acceptability of the test water would be its successful use in the culturing 

of L. plumulosus (with the control sediment).

10.2.5.2 Natural seawater or synthetic salt water can be used in the NAF/SBM 

testing program. Natural salt water should be obtained from an 

uncontaminated area known to support a healthy, reproducing population 

of L. plumulosus or similar sensitive species. Reconstituted salt water can 

be prepared by adding commercially available sea salt in specified 

quantities. Natural seawater should be filtered by passing through a 5 

micron filter before use. The reader is referred to ASTM E 1367-92 or E 

1367-99 for more information concerning test water.

10.3 Procedures

10.3.1 Mixing NAFs or SBMs with Control Sediment

EPA Method 1646 describes the method that should be used for amending control 

sediments with NAFs and SBMs. The control sediment should be sieved and 

homogenized before wet to dry weight ratio and density determinations are made and 

before NAFs are added to the control sediment. The following steps were given in 

EPA Method 1646 for mixing NAFs and SBMs with control sediments (parentheses 

were added here to provide additional information).

10.3.1.1 Determine the wet to dry weight ratio for the control sediment (three 

replicates of 30 g each has been used successfully)

10.3.1.2 Determine the density (g/ml) of the control sediment (three replicates of 

>25 ml is suitable for this purpose);

10.3.1.3 Determine the amount of NAF or SBM needed to obtain a desired test 

concentration; 
10-7

10.3.1.4 Determine the amount of wet sediment required;

10.3.1.5 Determine the amount of dry sediment in kilograms for each test 

concentration;

10.3.1.6 Determine the amount of NAF or SBM required to amend the control 

sediment at each test concentration;

10.3.1.7 Mix NAF or SBM with control sediment;

10.3.1.8 Test for homogeneity of NAF or SBM in sediment, and;

10.3.1.9 Mix sufficient quantities of NAF or SBM with control sediment for each 

treatment of amended or spiked sediment.

10.3.2 Estimate of Require NAF and SBM Required 

The six steps given above for NAFs can also be applied to SBMs, except that the 

third bullet in Step 3 requires a measurement of the density of the SBM. The density

of the SBM can then be used to estimate the quantity required for the desired test 

concentration. Refer to the formulas below for NAF and SBM calculations:

[10-1]

SBM Required (g) = [Conc. Desired (ml/kg)] x [Dry Weight Sediment (kg) x [SBM Density (g/ml)] [10-2]

See 40 CFR 435.13, 435.15, 435.43, and 435.45 for more information on this procedure.

10.3.3 Mixing Procedure

Mixing the NAF or SBM with the control sediment should be accomplished by

following these steps. The control sediment alone should also be subjected to the 

mixing procedure to ensure mixing has no effect on sediment toxicity.

10.3.3.1 Place appropriate amounts of weighed NAF or SBM into a stainless steel 

mixing bowl.

10.3.3.2 Tare the mixing bowl weight.

10.3.3.3 Add appropriate amount of control sediment.

10.3.3.4 Mix for 9 to15 minutes with a hand-held mixer equipped with stainless 

steel blades (e.g., KitchenAid Model KHM6).

10.3.3.5 As appropriate, test mixing homogeneity as described in Section 10.3.4.

10.3.4 Homogeneity of Mixing

NAF Required (g) =

[Conc. Desired (mg/kg)]

×

[Dry Weight Sediment (g)]

1000 g/kg 1000 mg/g
10-8

10.3.4.1 Tests for homogeneity of mixing should be performed, preferably in the 

procedure development phase (see 40 CFR 435.13, 435.15, 435.43, and 

435.45) by each laboratory performing NAF/SBM toxicity testing. This is 

to ensure that the NAF or SBM, which can be difficult to homogenize with 

control sediments, can be evenly mixed with the control sediment by each 

testing laboratory. 

10.3.4.2 EPA Method 1646 specifies that the coefficient of variation (CV) for a 

minimum of three replicate samples of the NAF/control sediment mixture 

must be less than 20%. Determinations of CV should be based on total 

petroleum hydrocarbon (TPH) content of the NAF or SBM as measured 

by EPA Methods 3550A and 8015M. If the initial CV is 20%, then the 

NAF/SBM-sediment mixture must be re-mixed and reanalyzed until the 

20% CV limit is achieved.

10.3.4.3 Homogeneity measurements should be made on the lowest and highest 

NAF concentrations for a given test. Laboratories should validate mixing 

efficiency via TPH measurements (as outlined above) of the low and high 

NAF concentrations. The homogeneity measurements should be made at 

least once per year.

10.4 Recommended Test Conditions

10.4.1 The recommended test conditions for conducting the 10-day or 96-hr sediment 

toxicity test with L. plumulosus are summarized in Table 3 and are consistent with 

methods presented in ASTM E 1367-92 and subsequent updates (E 1367-99). Tests 

should be conducted at 20±1 oC and 20±1‰ salinity with a 14h light; 10 h dark 

photoperiod at approximately 500-1,000 lux (or about 46 to 93 footcandles). Test 

chambers are 1-L glass containers with about a 10 cm inside diameter opening (or 

similar glass containers) that can contain about 150 ml sediment and 600 ml 

overlying water to achieve a 4:1 (v/v) water to sediment ratio. 

10.4.2 There are five (5) test concentrations plus a control for each NAF and SBM test. Five 

(5) replicates are included for the control sediment (E 1367-99) and for each test 

concentration.

10.4.3 The control sediment/test material mixture and test water should be added to test 

chambers the day before amphipods are added. This will allow for suspended 

particles to settle and allow time for equilibration of temperature and the sedimentwater interface. 

10.4.4 After the overnight equilibration period, amphipods are randomly distributed to each 

test chamber. Twenty amphipods are added to each replicate and there are five 

replicates per test treatment. 

10.4.5 Amphipods caught on the water surface can be pushed under with a glass rod. 

Individuals that have not burrowed within 5 to 10 minutes can be replaced, unless 

they are exhibiting an avoidance response. Amphipods are not removed at any time 

during the course of the toxicity test even if they appear dead. Test water is not 
10-9

renewed (i.e., static) and the amphipods are not fed during the exposure period. The 

toxicity test is terminated after 96 hours or 10 days for SBMs and NAFs respectively.

10.4.6 Temperature, salinity, pH, and dissolved oxygen (DO) should be monitored daily.

10.4.7 Ammonia should also be monitored in overlying water to ensure that the 

concentrations of this constituent do not exceed the tolerance range of the test species. 

For L. plumulosus, this is about 60 mg/L (as total ammonia) at pH 7.7 in 10-day tests 

(U.S. EPA, 1994). Ammonia has not been a problem in initial L. plumulosus 96-hr 

and 10-day tests with various NAFs.

Table 10-3. Conditions for conducting 96-hour NAF and 10-day SBM sediment toxicity 

tests with L. plumulosus. Conditions listed are consistent with test conditions specified in 

ASTM E 1367- 92 and subsequent updates (E 1367-99) unless otherwise noted

Parameter Conditions

Test Type Static whole sediment toxicity test

Temperature 20±1 oC

Salinity 20±1‰

Light Quality Wide-spectrum fluorescent or cool white lights

Illuminance 500-1,000 lux

Photoperiod 14h light: 10h dark *

Test Chamber 1-L glass beaker or jar

Sediment Volume 150 ml (2 cm depth)

Overlying water volume 600 ml (4:1 [v/v] water to sediment ratio)

Renewal of overlying water None

Size and life stage of 

amphipods 3-5 mm; immature and adult

Number of 

organisms/chamber 20

Number of test concentrations 5

Number of replicate 

chambers/treatment 5 in both controls and test treatments

Feeding None

Aeration Water in each test chamber should be

aerated throughout the test.

Overlying Water Clean natural or synthetic seawater

Overlying water quality Temperature, salinity, pH, and D.O.

daily; ammonia, as needed

Test duration 96 hours or 10 days

Endpoint Survival

Test acceptability Minimum mean control survival of 90% and satisfaction of 

criteria outlined in Table 4.

*Note: Although ASTM E 1367 specifies 16h light:8h dark, the photoperiod was changed to 14h light:10h dark to 

be consistent with the Mysidopsis bahia bioassay for drilling fluids (EPA Method 1619).
10-10

10.5 Biological Data 

10.5.1 Mortality is the endpoint for L. plumulosus at the end of the exposure period. At test 

termination, the contents of each test chamber (amphipods plus test sediment) are 

sieved through a 500 mesh screen to remove amphipods. 

10.5.2 Material retained on the screen should be rinsed into a sorting tray with clean salt 

water. 

10.5.3 The total numbers of live and dead amphipods should be recorded. 

10.5.4 Missing animals are presumed to have died and decomposed during the test and 

disintegrated. 

10.5.5 Amphipods should be counted alive if there are any signs of movement, such as a 

neuromuscular pleopod twitch (ASTM E 1367-99). Gentle prodding may be used to 

elicit movement.

10.6 Quality Control

10.6.1 Table 4 provides the acceptability requirements for the 10-day NAF and 96-hr SBM 

test. The primary acceptability requirement for NAF testing is as follows.

10.6.2 A toxicity test is unacceptable if more than a total of 10% of the control organisms 

die, or if the coefficient of variation (CV) of control survival is equal to or greater 

than 40%.

10.6.3 If this acceptability requirement is not met, then the data should be discarded and the 

experiment repeated. 

10.6.4 If this requirement is met, then the other acceptability requirements in Table 4 should 

be reviewed and a determination made as to the acceptability of the data.
10-11

Table 10-4. Test acceptability requirements for 10-day NAF and 96-hr SBM tests with L. 

plumulosus. Requirements listed are consistent with those specified in ASTM E 1367-92 

and subsequent updates (E 1367-99)

Ten-day NAF and 96-hr SBM toxicity tests should usually be considered unacceptable if one or 

more of the following occurred:*

 A 10-day NAF and 96-hr SBM toxicity tests are unacceptable if more than a total of 10% 

of the control organisms die, or if the coefficient of variation (CV) of control survival is 

equal to or greater than 40%.

 All test chambers were not identical.

 Test organisms were not randomly or impartially distributed to test chambers.

 Required reference standard was not included in the test.

 All test animals were not from the same population, were not all of the same species, or 

were not of acceptable quality.

 Amphipods from a wild population were maintained in the laboratory for more than two 

weeks, unless the effects of prolonged maintenance in the laboratory has been shown to 

have no significant effect on sensitivity.

 The test organisms were not acclimated at the test temperature and salinity at least 48 

hours before they were placed in the test chambers.

 Temperature and dissolved oxygen concentrations were not measured

*Note: These guidelines are not identical to those listed ASTM E 1367 in part because some acceptability guidelines 

listed in ASTM E1367-92 are not applicable or practical for the NAF/SBM toxicity testing program.

10.7 Reference Tests 

10.7.1 A single toxicity test will be used to determine satisfactory laboratory performance 

and to determine whether an NAF or SBM can be discharged as it adheres to drill 

cuttings. The reference toxicant for the NAF test will be either a C16-C18 internal 

olefin reference standard or a C12-C14 or C8 ester. 

10.7.2 The reference toxicant for the SBM testing program will be a C16-C18 internal olefin 

SBM which has also been specified for determining pass/fail for SBMs. The C16-C18

internal olefin SBM is a 65/35 blend, proportioned by mass, of hexadecene and 

octadecene, respectively (see 40 CFR 435.11(rr). 

10.7.3 These reference toxicity tests will be conducted in conjunction with all NAF or SBM 

tests to discern possible changes in the condition of the L. plumulosus population used 

in testing. The reference toxicant test must be conducted concurrently with each 

sample or batch of samples and at a minimum should be conducted at least monthly. 

Control charts of this reference standard should be maintained to perform statistical 

analyses, help understand the inherent variability in the reference test, and for longterm quality control. Test conditions for the reference test should follow the 

experimental conditions presented in Table 3.

10.7.4 The reference toxicant test should be performed concurrently-and under the same 

conditions as the NAF or SBM test. 
10-12

10.7.5 The reference toxicant test should be conducted so that control limits (typically set at 

2 standard deviations) can be established (U.S. EPA, 1994). 

10.7.6 If the reference test LC50 falls outside of this range of control limits generated on the 

most recent test data points, then the sensitivity of L. plumulosus and the credibility of 

the test results are considered suspect. In this case, the test procedure should be 

examined and the test repeated with a different batch of amphipods. A sediment test 

should not automatically be judged unacceptable if the reference test LC50 falls 

outside the expected range or if the control in the reference toxicity test exceeds 10%. 

The width of the control limits and all performance criteria listed in Table 4 should be 

considered when determining the acceptability of a given NAF or SBM test.

10.8 Interpretation of Result

10.8.1 Procedures presented in this test method are used to calculate point estimates, or LC50

values. The LC50 value and 95% confidence limits of the NAF tests should be 

calculated on the basis of milligrams of NAF per kg dry control sediment (mg/kg) and 

amphipod mortality. 

10.8.2 The LC50 value and 95% confidence limits of the NAF tests should be calculated on 

the basis of milliliters of NAF per kg dry control sediment (ml/kg) and amphipod 

mortality. 

10.8.3 A variety of methods can be used to calculate an LC50 value and its 95% confidence 

limits, including probit, moving average, trimmed Spearman-Karber and LitchfieldWilcoxon methods (ASTM E 1367-99). The method used should take into account 

the number of partial kills, the number of test chambers per treatment (5), and the 

number of amphipods per test chamber (20).

10.8.4 The only NAF-drill cuttings that can be discharge are cuttings coated with NAFs that 

are as toxic or less toxic, but not more toxic, than the reference NAF (C16-C18 internal 

olefin or C12-C14 or C8 ester) as measured by LC50. This limitation is expressed in 40 

CFR 435.13, 435.15, 435.43, and 435.45.

[10-3]

10.8.5 The only SBM-drill cuttings that can be discharge are cuttings coated with SBMs that 

are as toxic or less toxic, but not more toxic, than the reference SBM (C16-C18 internal 

olefin or C12-C14 or C8 ester) as measured by LC50. This limitation is expressed in 40 

CFR 435.13, 435.15, 435.43, and 435.45.

[10-4]

10-day LC50 Reference Material ≤ 1.0 10-day LC50 NAF

96-hr LC50 Reference Drilling Fluid ≤ 1.0 10-day LC50 SBM
10-13

10.8.6 The NAF or SBM data should be interpreted by comparing to the reference toxicant 

test LC50 value and whether it exceeds the limitations in 40 CFR 435.13, 435.15, 

435.43, and 435.45.

10.9 References

10.9.1 American Society for Testing and Materials (ASTM). 1997. Standard Guide for 

Conducting 10- day Static Sediment Toxicity Tests with Marine and Estuarine 

Amphipods. E 1367-92. Annual Book of ASTM Standards, Vol. 11.05.

10.9.2 American Society for Testing and Materials (ASTM). 2000. Standard Guide for 

Conducting 10- day Static Sediment Toxicity Tests with Marine and Estuarine 

Amphipods. E 1367-99. Annual Book of ASTM Standards, Vol. 11.05.

10.9.3 Emery, V.L., D.W. Moore, B.R. Gray, B.M. Duke, A.B. Gibson, R.B. Wright and 

J.D. Farrar. 1997. Development of a chronic sublethal sediment bioassay using the 

estuarine amphipod Leptocheirus plumulosus (Shoemaker). Environ. Toxicol. Chem. 

16:1912-1920.

10.9.4 McGee, B.L. D.J. Fisher, L.T. Yonkos, G.P. Ziegler and S. Turley. 1999. Assessment 

of sediment contamination, acute toxicity, and population viability of the estuarine 

amphipod Leptocheirus plumulosus in Baltimore Harbor, Maryland, USA. Environ. 

Toxicol. Chem. 18:2151-2160.

10.9.5 Schlekat, C.E., B.L. McGee and E. Reinharz. 1992. Testing sediment toxicity in 

Chesapeake Bay with the amphipod Leptocheirus plumulosus: an evaluation. 

Environ. Toxicol. Chem. 11:225-236.

10.9.6 Suedel, B.C. and J.H. Rodgers, Jr. 1991. Variability of bottom sediment 

characteristics of the continental United States. Water Res. Bull. 27:101-109.

10.9.7 U.S. EPA, 1994. Methods for Assessing the Toxicity of Sediment-associated 

Contaminants with Estuarine and Marine Amphipods. EPA/600/R-94/025. USEPA 

Office of Research and Development, June 1994.

10.9.8 U.S. EPA, 2001. Effluent Limitations Guidelines and New Source Performance 

Standards for the Oil and Gas Extraction Point Source Category; OMB Approval 

Under the Paperwork Reduction Act: Technical Amendment, Federal Register, 

Volume 66, Page 6849-6919, January 22nd.