The
   DENTIFICATION
and
  MEASUREMENT
 of
  CHLORINATED
  HYDROCARBON
  PESTICIDES
  in
  SURFACE WATERS
  U.S. DEPARTMENT OF THE INTERIOR
  Federal Water Pollution Control Administration

-------
The
   IDENTIFICATION
and
   MEASUREMENT
 of
   CHLORINATED
   HYDROCARBON
   PESTICIDES
  in
   SURFACE WATERS
By  A.W. BREIDENBACH
   JJ. LICHTENBERG
   C.F. HENKE
   DJ. SMITH
   J.W. EICHELBERGER, JR.
   H. STIERLI
   Water Quality Activities
   Division of Pollution Surveillance
   U.S. DEPARTMENT OF THE INTERIOR
   Federal Water Pollution Control Administration
   Washington, D.C. 20242

-------
                       A NOTE ABOUT THE  AUTHORS

         Dr. Breidenbach is Officer in Charge of the Water Pollution Surveillance Sys-
tem and Mr. Lichtenberg is supervisor of the Organic Chemistry Laboratory.
         Mrs.  Smith and Mr. Eichelberger are chemists working on the identification
and quantification of organic compounds.  Mr. Henke  supervises the extraction  and
separation activities.  Mr. Stierli, a registered professional engineer, serves as Chief of
Engineering and Field  Operations for  the Activities.
                            ACKNOWLEDGEMENT

         The authors wish to express their sincere appreciation to Dr. F. K. Kawahara,
Consultant, Organic Chemistry, for  his valuable assistance in the review of the several
procedures and the editing of the manuscript.  The excellent work of Mrs. D. Platt in
preparing the final copy is gratefully acknowledged.

-------
PREFACE
        In September 1964 when this manual was first issued as an informal publi-
cation (PHS Publication #1241), it was recognized  that  anticipated progress in
methods  development would require periodic revision of the text;  This issuance up-
dates the previous .effort  and describes  the • methods  currently  employed  in the
Surveillance  System  laboratories  of  the Federal   Water  Pollution  Control  Ad-
ministration.
        Organic  chemicals, as a group,  have presented a  special challenge to the
laboratory  because of the  many  thousands of such  chemicals in use and the many
complex mixtures of wastes produced in their manufacture. Specific identification and
measurement of one class  of organics, the chlorinated  hydrocarbon  pesticides, to a
sensitivity of one microgram per  liter or below is of particular concern.
        The  carbon adsorption method, developed over a decade ago, has been effec-
tively employed in pesticide pollution studies. This method was pioneered and developed
by a team of scientists at the Robert A. Taft Sanitary Engineering Center of the Public
Health Service: F. M. Middleton, M. Ettinger, A. Rosen,  G. Walton and H. Braus.
While it is essentially a qualitative screening and continuous sampling technique when
used on untreated surface  waters, the method provides minimum quantitative values
for measurement of specific substances.  Most  significant, the method has proved to be
very useful for  obtaining samples  large enough for corroborative infrared and chroma-
tographic identifications  at  low concentration levels.
        Chromatography and chromatographic instrumentation have made possible the
development and application of additional techniques by the Federal Water Pollution
Control Administration's surveillance system laboratories.  These  newer techniques,
applied to carbon adsorption extracts as  well as discrete  water samples, have been used
to provide definitive identification and measurement of chlorinated hydrocarbon pesti-
cides in surface waters. The rapid progress being made in methods research will surely
result in  continued modifications, improvements, and additions.
        Further developmental work is going forward within the laboratory elements
of the FWPCA and in other Federal  agencies which have responsibilities associated
with pesticides  in  the various segments of the environment.  The Federal Committee
on Pest  Control provides guidance and  coordination  to interagency efforts in this
problem area.


                                            RICHARD S. GREEN, Acting Chief
                                                Division of Pollution Surveillance
                                                          Federal Water Pollution
                                                           Control Administration
                                       m

-------
                     CONTENTS
                                                   PAGE
 PREFACE 	:	     iii
 I. INTRODUCTION  	     1
 A. BACKGROUND	     1
 1. Federal Water Pollution Control Administration Water
   Pollution Surveillance System	     1
 2. Organic Pollution	     1
 B. CARBON  ADSORPTION SAMPLING 	     3
 C. BOTTLED SAMPLES 	     3
. 1. Water  Samples	     3
 2. Bottom Samples	     4
 II. SAMPLE  COLLECTION	     5
 A. THE CARBON ADSORPTION METHOD (CAM) ....     5
 1. Preparation of the Carbon Adsorption Cartridge	     5
 2. Precautions Necessary to Prevent Accidental
   Contamination of Carbon	     6
 B. DISCRETE BOTTLED SAMPLES	     6
 1. Water  Samples	     6
 2. Bottom Samples	     6
 3. Preparation of Container	     7
 III.  PREPARATION OF SAMPLES PRELIMINARY TO
     GAS  CHROMATOGRAPHIC ANALYSIS  	     8
 A. CARBON  ADSORPTION SAMPLES	     8
 1. Treatment of  Carbon  	     8
   a. Drying  the Carbon	     8
   b. Extraction of the Carbon	     8
 2. Preliminary Separation of CCE	     12
   a. Procedure for General Organic Analysis	     12
   b. Column Chromatographic Separation of CCE
      (Alternate Procedure) 	     15
   c. Thin Layer Chromatographic (TLC) Separation of
      Pesticides from Aromatic Fraction of CCE	     17

-------
                                                    PAGE
B. DISCRETE BOTTLED SAMPLES	    22
1.  Extraction of Pesticides	    22
   a. Extraction from Water 	    22
   b. Extraction of Bottom Samples  	    25
2.  Concentration of Extract	    25
   a. Water  Sample  	    25
   b. Bottom Sample 	    25
3.  Thin Layer Chromatography 	    27
IV. DETERMINATIVE STEPS  	    28
A. GAS CHROMATOGRAPHY	    28
1.  Application of Electron Capture Gas Chromatography  . .    28
2.  Application of Microcoulometric Titration Gas
   Chromatography  	    30
3.  Calculations  	    30
4.  Column Packings	    31
5.  Column Conditioning	    32
B. INFRARED SPECTROPHOTOMETRY	    32
V. CONTROL OF INTERFERENCES	    33
A. SOLVENT INTERFERENCES 	    33
1.  Chloroform	    33
2.  Hexane-Benzene	    33
3.  Hexane-Acetone 	    34
4.  Carbon Tetrachloride and Acetone	    34
B. CARBON INTERFERENCES 	    34
1.  Carbon Blank	    34
C. OTHER SOURCES OF INTERFERENCE	    34
D. INTERPRETATION  	    35
VI. SENSITIVITY AND SPECIFICITY	    36
A. SENSITIVITY  	    36
1.  Carbon Adsorption Extracts Examined by Electron
   Capture Gas Chromatography  	    36
2.  Carbon Adsorption Extracts Examined by Micro-
   coulometric Titration Gas Chromatography 	    37
3.  Bottled Sample  Extracts Examined by Electron Capture
   Gas Chromatography 	    37
4.  Bottled Sample  Extracts Examined by Microcoulometric
   Titration  Gas Chromatography 	    37

                            vi

-------
                                                    PAGE

B. SPECIFICITY  	    37
1.  Carbon Adsorption Samples  	    37
2.  Bottled Samples  	    37

APPENDIX ONE	    38
Engineering Aspects  of Sampling by the Carbon
  Adsorption Method

APPENDIX TWO  	    51
Chromatograms, Sample Calibration Curves, Infrared
  Spectra, and Structural Formulae

APPENDIX THREE 	    61
Equipment, Solvents and Reagents

APPENDIX FOUR 	    64
General Composition of Carbon Chloroform and Carbon
  Alcohol Extracts

APPENDIX FIVE	    67
Glossary

REFERENCES 	    67
                    LIST OF TABLES

                                                    PAGE
TABLE 1  	    58

Rf Values of Pesticides Developed with CCU on
Silica Gel-G Thin Layer Plate

TABLE 2  	    59
Gas Chromatographic Retention Data

TABLE 3  	    60
Some Column Packings Used for Gas Chromatographic
Analysis of Chlorinated Hydrocarbon Pesticides
                           vn

-------
                 LIST OF ILLUSTRATIONS

FIGURE                    TITLE                      PAGE

   1  FWPCA Water Pollution Surveillance System
        Sampling Stations  	     2
   2  Screw-cap (Teflon-lined), Glass Sample Containers
        and Expanded Polystyrene Cartons	     7

   3  Laboratory Data Card	     9

   4  Carbon Drying Oven	    10

   5  Soxhlet Extractors	    11

   6  Removal of Residual Chloroform from Carbon	    11

   7  Flow Scheme for Solubility Separation of CCE	    13

   8  Flow Scheme  for Chromatographic Separation  of
        Neutrals  	    15

   9  Infrared  Spectrum  of  Aromatic  Fraction  of CCE
        Sample Supporting Chromatographic  Identification
      of DDT  	    16

  10  Infrared  Spectrum  of  Aromatic  Fraction  of CCE
        Sample. Supporting Chromatographic  Identification
        of Dieldrin  	    16

  11  Spotting of TLC Plate	    18

  12  Diagram  of Designation of Sections in the Cleanup
        and Separation of CCE-Aromatics  on Silica Gel
        Layers  	    19

  13  Photograph of a Developed Thin Layer Plate	    20

  14  Silica Gel Collection Assembly	    21

  15  Semi-automatic Liquid-Liquid Extractor	    23

  16  Solvent Recovery Apparatus	    24

  17  Soxhlet Extraction of Bottom Samples	    26

  18  Calculation of Peak Area	    29

  19  Carbon Adsorption Column and Shipping Container ..    39

  20  Details of Sand Prefilter	    40

  21  Schematic Diagram of an Installation with Manual
        Backwash  	    41

  22  Carbon Adsorption Unit  Model H20-M1C with Sand
        Prefilter and Automatic  Backwash  	:	    42

  23  Carbon Adsorption Unit Model H20-M2C with Pre-
        settling Tank and Auxiliary Equipment in Shelter . .    43
                             vin

-------
FIGURE                     TITLE                       PAGE

  24  Schematic Flow Diagram with Sampling Procedure for
        Organic Sampler Models H20-M1C and H20-M2C    44
  25  Equipment Installed  in Field Test Station for Field
        Evaluation of Low Flow  Rate Samples  in  Com-
        parison with Conventional  Sampling Apparatus ...     46
  26  Low Flow Rate Organics Sampler, Model LF-1  	     47
  27  Low Flow Rate Organics Sampler, Model LF-2	     48

  28  EC Gas Chromatogram of Standard Pesticides in TLC
        Section II (heptachlor epoxide, dieldrin, endrin)  ..     51
      EC Gas Chromatogram of Standard Pesticides in TLC
        Section III (lindane, y-chlordane, DDD)  . ..	     51
      EC Gas Chromatogram of Standard Pesticides in TLC
        Section IV (heptachlor, aldrin, DDE, DDT)  	     51
      Sample Calibration Curve for Dieldrin (ECGC) ....     51

  29  MCT Gas  Chromatogram of Standard Pesticides in
        TLC  Section II  (heptachlor  epoxide,  dieldrin,
        endrin)  	     52
      MCT Gas  Chromatogram of Standard Pesticides in
        TLC Section III  (lindane, y-chlordane, DDD) 	     52
      MCT Gas  Chromatogram of Standard Pesticides in
        TLC Section IV (heptachlor, aldrin, DDE, DDT) . .     52
      Sample Calibration Curve for Endrin (MCTGC) 	     52
  30  IR Spectrum of Standard Dieldrin in Mineral Oil Mull    53
  31  IR Spectrum of Standard Endrin in Mineral Oil Mull     53
  32  IR Spectrum of Standard Lindane in Mineral Oil Mull    54
  33  IR Spectrum of Standard DDD in Mineral Oil Mull . .     54
  34  IR Spectrum of Standard Heptachlor in Mineral Oil
        Mull  	     55

  35  IR Spectrum of Standard Aldrin in Mineral Oil Mull     55
  36  IR Spectrum of Standard DDE in Mineral Oil Mull . .     56
  37  IR Spectrum of Standard DDT in Mineral Oil Mull . .     56
  38  Structural Formulae of Nine Chlorinated Hydrocarbon
        Pesticides	     57
                             IX

-------
I.  INTRODUCTION
A.   BACKGROUND


1.  Federal Water Pollution  Control Administration Surveillance System

        The surveillance  system l was established under the Public Health Service
in October 1957 to implement that part of Public Law 660, as later amended, wherein
the Secretary of the Department of Health, Education, and Welfare was authorized
to collect and disseminate basic data on chemical,  physical,  and  biological water
quality  insofar as  such data relate to water pollution, prevention, and  control.  The
system was expanded at the rate of 25 stations per year until it reached a total of 122
stations in September  1962. Six sampling stations were added during the following
year.  Two sampling stations were  established  in  1964 in the lower Mississippi main
stem and one at Morgan City, Louisiana  on the Atchafalaya. There are now a total
of 131 sampling stations.  Figure 1  shows the location of the stations.
        Participants in the system include more than 176 state, local, and Federal
water, sewage or other public utilities, health departments, industries, universities, state
water pollution agencies, and  resident engineers of Federal reservoirs.  Active local
participation is important in this operation.  The state  and local  agencies perform
most of the conventional  chemical  analyses and  collect  water samples  for the more
complex examinations.  The FWPCA performs the more complex determinations at
its Cincinnati laboratories  and  makes the results available to the various participants.
The program as a whole is designed to assemble, examine, and interpret the facts which
enable water pollution  control  agencies and others concerned to determine the scope
and character of problems  to be solved.
        The analytical work of the surveillance system is devoted to characterization
of surface water samples in six broad  disciplinary areas.  These are biological, micro-
biological  and particulate  matter,  radiological, general chemical as well as physical
properties and synthetic organic chemicals. Frequency of collection of the various dis-
crete samples varies from  several times per hour with automatic  field instrumentation
to once per month, depending on the type  and purpose of the sample.


2.  Organic Pollution

        A very large variety of organic  pollutants is known to be  present in river
water. These substances, present in small concentrations, may be carried to the stream
 Formerly the National Water Quality Network.

-------
             FVVPCA  Water Pollution  Surveillance System
                                  SAMPLING STATIONS
2 Stations in Alaska not shown
 Anchorage and Fairbanks
             Figure  1. FWPCA Water Pollution Surveillance System Sampling Stations

-------
in runoff, in domestic sewage or in industrial wastes.  They are carried in solution and
adsorbed to suspended solids. The 5-day biochemical oxygen demand (BOD), chemi-
cal oxygen demand (COD), nitrogen analyses and total carbon have been used success-
fully to aid in describing the degree and type of organic pollution as well as estimating
oxidizability in the stream  environment.  These tests do not, however, serve as tools
to identify  and measure the specific organic compounds  which are present in polluted
water. The rapid and economical measurement of microgram and nanogram quantities
of organic pollutants in water has been extremely  difficult and  impractical until very
recently. Indeed, such minute quantities of specific substances, intermixed with a large
variety of other interfering organic substances, have presented an extremely challenging
and enigmatic analytical problem. Until recently,  identification and measurement of
most organic compounds in water in the parts per billion sensitivity range, has required
an extremely large sample.  Most methods, insofar as they have been  developed, are
not yet sufficiently sensitive to be of  use with smaller samples.
        The need  for larger samples for exploratory work  in characterizing organic
pollutants was recognized over a decade ago and stimulated  several  years of research
which ultimately produced the carbon adsorption method (1)  (2) (3) (4) (5) (6) (7).
B.  CARBON  ADSORPTION SAMPLES

        This method, described in detail herein, uses the adsorptive capacity of acti-
vated carbon to concentrate organic materials  from large water samples, measuring
from 300 to 5000 gallons.  This  large sample permits corroborative identification  by
several analytical methods which can provide highly defensible identifications of specific
substances. It must be noted, however, that the  concentration values obtained for spe-
cific substances with this method must be considered as  minimum values.  First, the
efficiency of the adsorption  on and the desorption from carbon cannot be expected to be
100 percent for all compounds under widely varying physical and physico-chemical con-
ditions in the water being sampled. Studies have shown recently, for example, that the
adsorption of organics in streams on carbon is most efficient at flow rates and throughput
volumes less than those which have been employed previously  (8) (9)  (10)  (11)  (12).
Secondly, until a means is available to gather organics adsorbed on the suspended solids
as well as the organics in solution from the large sample, the determined concentration
values  (e.g., microgram per liter of  water) must  be  considered  low.  The  increased
yields per unit volume of water  from low flow equipment are primarily due to the
longer contact time. However, they may result .in part from trapping of and subse-
quent desorption from some of the suspended solids. Pesticides have been identified in
carbon adsorption samples in the past (13)  (14) (15)  (16)  (17).
C.   BOTTLED  SAMPLES

1.  Water Samples

        Water grab samples of one liter to one gallon are useful. The grab sample,
properly taken, contains water in which organics are dissolved as well as suspended
solids on which organics are adsorbed. The absolute weight of the organic material is

-------
so small in  most grab samples as to restrict the approach used for identification to
noninfrared analysis.  However, if the organic substance present can be detected, it can
usually be measured at very low levels.  Thus, in the microgram per liter concentra-
tion range,  carbon adsorption samples provide enough material so  that the potential
for corroborated identification exists. Grab samples are  most useful  for rapid  and
highly sensitive measurement and, in addition, supply further data on which the iden-
tification  can be based. A number of workers have identified pesticides in water grab
samples (18) (19) (20) (21).


2.  Bottom  Samples

        Pesticides have been identified in  silt (22)  and  soils (23) (24) (25)  (26).
The water in a stream is closely related to the solids suspended in it, as well as  the solids
deposited  on the bottom. Solids accumulated on the bed of a stream may contain larger
quantities of organics than the water above.  Extensive studies on the Lower Mississippi
River have  shown this to be true in the case of certain chlorinated pesticides.  There-
fore, it is  equally important to analyze the  bottom samples as  well when assessing the
degree of  a pollution problem.

-------
II.  SAMPLE  COLLECTION
        Carbon adsorption samples and discrete one-liter samples  are taken for the
identification and measurement of organic substances  in  surface water.2  These sam-
pling approaches have been combined with sensitive thin layer chromatographic  and
gas chromatographic methods for  pesticide analysis. The details of the sampling tech-
niques  employed in the laboratories of the FWPCA  surveillance system are outlined
below.
A.   THE CARBON  ADSORPTION  METHOD  (CAM)

        This technique, developed in 1951 (1), was applied to raw surface waters in
a pilot study in 1956 and  has been in routine use in  the surveillance system since
1957. Since that time  the sample collection aspects of the technique have undergone
considerable refinement. Pertinent details of the sampling equipment are included in
Appendix One.


1.  Preparation of the Carbon Adsorption  Cartridge (CAC)

        The carbon adsorption cartridge consists of a Pyrex glass pipe three inches in
diameter and 18 inches in  length packed with  two types of carbon.  To pack the ver-
tically oriented cylinder are  added successively  4.5 inches  of 4 x  10 mesh carbon,8
nine inches of 30 mesh carbon,4 and an additional 4.5  inches of 4 x 10 mesh carbon.
The cylinder is packed full but not tightly.  The coarse  carbon at each end of the
cartridge aids in preventing clogging by mud  and silt  from  turbid waters. However,
cartridges employing all fine  (C-190) carbon at low flow rates and reduced through-
put volumes are being used successfully (see Appendix One). The cartridge is shipped
to the field station and installed in the appropriate sampling system.
        Sample volumes of 300 and 5000 gallons (1)  (27)  (28)  of water taken at
rates of 0.03 and 0.5 gpm have  been used successfully; however, sampling efficiency
can  be increased by the use  of smaller volumes  and  lower flow rates.  Efficiency is
further increased with the  low  flow rate system  using  a  column containing  only 30
mesh carbon. The reproducibility  (29) and effect of the variables of total through-
put and rate of flow through  the carbon (10) have been the subject of intense study
- Although these methods are employed currently for untreated surface water the methods are also
applicable to ground water and treated water.
:! Cliff Char 4 x 10 mesh (Cliffs Dow Chemical Co., Marquette, Mich.)
4 Nuchar C-190 (West Virginia Pulp and Paper Co., New York, N.Y.)

-------
by Booth.  A sampling system for lower flow rates and lower throughput volumes was
designed by  Castelli  and Booth (30).  Field tests show that sampling efficiency  is
greatly improved  under  these conditions  (31) (see Appendix One).
        After the desired quantity of water is sampled, the carbon cartridge is dis-
connected and returned to the laboratory for analysis  (see Figure 19, Appendix One).
2.  Precautions Necessary to Prevent Accidental Contamination of Carbon

         The affinity of carbon for organic substances requires that supplies of carbon
be protected from extraneous sources of contamination. For example, carbon can ad-
sorb organic substances such as paint vehicles and  insecticides used for pest control
from the air.  Therefore, the carbon is stored and processed  in an area adequately
protected from  such  sources  of  contamination.  As an  additional  precaution, the
ventilating,  heating, and air conditioning systems for the laboratories  in which  car-
bon adsorption samples are processed  are completely isolated from all other labora-
tories.  All carbon is obtained from the manufacturer in sealed metal drums. Obviously,
spraying with pest control chemicals is not permitted in  these  areas.  Carbon blank
determinations supplement these precautions (see  Section V).
B.  DISCRETE  BOTTLED  SAMPLES
1.  Water Samples

        Approximately one liter (940 ml) of water is collected in each, of two wide-
mouthed glass bottles equipped with screw caps fitted with Teflon liners.  The bottle
should be filled to about y2 inch from the top. The mouth of the bottle must be wiped
clean before securing the cap to prevent leaking.  These  two bottles  represent one
sample.  Plastic bottles  (polyethylene) are not used because traces of  plasticizer are
leached from the plastic by the water and can be a source of analytical interference.
Moreover, organics from the water are adsorbed on the plastic.  It has been suggested
that high grade Teflon (Nalgene) bottles may be satisfactory for this use; however, the
cost is prohibitive at present.  Many investigators avoid the use of glass sample bottles
because breakage in shipment frequently causes loss of sample.  This is overcome by
the use of  relatively  inexpensive,  expanded, polystyrene  foam  shipping  containers
molded to fit the bottle (see  Figure 2).
2.  Bottom  Samples

        Samples may be collected from the stream bottom with a St. Anthony Falls
type device (32).  The sample is placed in a 1-quart bottle such as  described in B.I.
above.  The bottle should  be about  V?  filled.  The samples  may be coarse or fine
gravel, sand, silt, or clay.

-------
Figure 2.  Screw-cap  (Teflon-lined),  Glass Sample Containers and Expanded
                              Polystyrene Cartons
3.  Preparation of the Container

        Bottles are  rinsed successively with chromate cleaning solution, running tap
water, distilled water,  and finally several times with redistilled solvent (e.g., acetone,
hexane, petroleum ether,  chloroform).  Caps and liners are washed with  detergent.
rinsed with tap water, distilled water and solvent.

-------
      PREPARATION  OF  SAMPLES  PRELIMINARY TO
      GAS CHROMATOGRAPHIC  ANALYSIS
        After a carbon adsorption or grab  sample is received, it is  logged and  all
pertinent data (source, date sampled, date received, quantity of water sampled) are
recorded (see Figure 3).
A.   CARBON  ADSORPTION SAMPLES

1.  Treatment of Carbon

a. Drying the Carbon

        The carbon is dried by spreading it on stainless  steel trays in  an oven at
40°C. for  about  two  days5-6  (see Figure 4).  If there is a backlog of dried carbon
samples on hand they are sealed in solvent-rinsed, one-gallon, wide-mouthed tin cans
and held for further treatment.

b. Extraction of  the Carbon
        Large scale  Soxhlet  extractors  are used to extract the  dried carbon.  The
quantity of carbon used in the sampling cartridge is accommodated by the extractors
(see Figure 5).
(1) Packing the  extractors
        To prevent carbon fines from passing into the boiling flask, the bottom of the
extractor is packed with about three inches of pre-extracted glass wool.7  The  wool
is wetted with chloroform. The dried carbon sample is added and packed by tamping so
that it just fits the extractor.  If carbon is packed  too tightly,  siphoning will be se-
verely hindered.  The  frequency of siphoning  is controlled  at 2 cylinder volumes per
hour. Siphoning is not always automatic  and application of  compressed air to the vent
of the extractor is sometimes necessary.
(2) Chloroform extraction
        The Soxhlet  is filled with  redistilled chloroform and siphoned over twice.
More chloroform is added, if necessary,  and the sample is  extracted  continuously for
•"' Copper or brass trays may also be used. Galvanized metals or aluminum react with wet carbon.
Metal coated with high quality Teflon has also been suggested.
n The air circulated through the oven is prefiltered through carbon to prevent contamination from
the atmosphere.
7 Oily organic substances are  first removed from glass wool by extraction with chloroform.

                                      8

-------
35 hours.8 After the extraction  is completed, the bulk of the chloroform is siphoned
and  blown over into  the  boiling flask.  The  flask is removed  from the  system, the
extract concentrated  to about 250 ml by distillation,9 and filtered  through  solvent-
washed filter paper  into  a 300-ml Erlenmeyer flask. The  solvent is evaporated  to
approximately 20 ml  on a steam bath with a jet of clean, dry  air.10  The contents  of
the flask  are transferred  to a tared glass vial and  the remaining  solvent evaporated
at room temperature in  a  hood without a jet of air. The carbon chloroform extract
(CCE) is judged dry when the chloroform odor can no  longer be  detected.11  The
weight of the residue is obtained.
8 Longer extraction times may be used but 35 hours (24 for ethanol) is considered optimum. Booth
(19) has confirmed this point.
9 Two-zone Glas-Col heating mantles are used  to prevent overheating and scorching of the sample.
10 First the compressed air is cleaned and dried by directing it through  a bed containing carbon and
a drying agent such as calcium chloride.
11 A trace of chloroform is retained by the CCE.  This  is, however, insignificant in most samples
and no correction is necessary. In very large fluid samples correction may be necessary and can
be accomplished using a procedure developed by Mashni  (33).  Unfortunately, it  is not practical
to  do this on a routine  basis for large numbers of samples.
       STANDARD ORGANIC ANALYSIS OF
       CARBON ADSORPTION SAMPLES
 STATION NO.	
 SAMPLE NO.	_
 DATE	TO	
 RECEIVED	
 FLOW IN GALLONS.
  SOURCE	
  LOCATION.
                                                   WEIGHT OF SAMPLES IN GRAMS
                                              CHLOROFORM EXTRACT     ALCOHOL EXTRACT
    .TYPE OF WATER -
.COMPOSITE.
               .QUARTER.
                      ..LITERS.
             EXTRACTION DATA
EXTRACT
CHLOROFORM
ALCOHOL
TOTAL
GRAMS



P. P. B.



PER CENT



DATE EXTRACTED



    SEPARATION OF CHLOROFORM EXTRACT
 REMARKS:
 SEC L6?a
 (6-41.)
SOLUBILITY SEPARATION
ETHER INSOLUBLES
WATER SOLUBLES
NEUTRALS
ALIPHATICS
AROMATICS
OXYS
LOSS
TOTAL
WEAK ACIDS
STRONG ACIDS
BASES
LOSS
TOTAL
P. P. B.








NEUTRALS






PER CENT








NEUTRALS









SOLUBILITY SEPARATION
CHLOROFORM EXTRACT



ETHER INSOLUBLES WATER SOLUBLES



WEAK AGIOS STRONG ACIDS



BASES NEUTRALS






CHROMATOGRAPHU
SEPARATION
NEUTRAL FRACTION



ALIPHATICS



AROMATICS



OXYS



                   MOST
                                                             Bid
                         Figure 3.  Laboratory Data Card

-------
                         Figure 4.  Carbon  Drying  Oven
(3) Ethanol extraction
         This step is not used in routine pesticide  analysis. The residual chloroform
is  removed  from the carbon by  blowing pre-cleaned warm  air through the carbon (in
place in the Soxhlet)  and exhausting chloroform vapors through the hood.  In  order
to do this the Soxhlet  is removed  from the hood and the carbon shaken loose to facili-
tate movement of  air through it.  The Soxhlet, still containing the carbon, is returned
to the hood with the glass plate cover removed.  A  hose from a heated air  manifold
(approximately 60° C.) is attached to the bottom of the siphon tube  and the  air i?
blown up through the  carbon  for three to four hours  or  until it is dry  (see Figure
6). Alternate methods,12'ia may be employed but this procedure has proved much less
hazardous,  consumes  less  time  and requires less supervision.  Ethanol (95%) is added
to the dried carbon and the extraction is carried out in the manner described for tilt-
chloroform  step.  Extraction is  terminated after 24 hours.8
         Concentration  of  the carbon  alcohol extract (CAE) is begun as described for
the chloroform extract. However,  the drying, started on a steam bath with a jet of air,
12 The carbon is removed from the Soxhlet and dried in the oven as in III, A, 1, a. This proce-
dure requires about 48 hours.  Adequate ventilation is required to remove hazardous  chloroform
vapors.
18 The residual chloroform  may be leached from the carbon by pouring alcohol over it. Proceed
as  follows: Siphon twice and distill until 68° C. is reached. Repeat a second and third time, distill
to  77° C. and begin extraction. Add alcohol if necessary.  The distillate  (68° to 77° C.) may bf-
used for the  initial leaching of the carbon in succeeding extractions.  This procedure requires about
4 hours.

                                         10

-------
           Figure 5. Soxhlet Extractors
Figure 6.  Removal of Residual Chloroform from Carbon




                         11

-------
is continued  in an oven  at 75° C. until weight change of successive weighings at 72-
hour intervals is less than \%.
2.  Preliminary Separation of CCE (27) (34) (35) (36) (37)

         The procedure described under paragraph 2a is employed when all classes of
compounds  are of interest. When only chlorinated hydrocarbon pesticide information
is sought, the procedure 2b is used.

         The separation techniques and analytical procedures described below are car-
ried out as quantitatively as possible.  Careful  attention  is directed  at  details of
quantitative transfer,  controlled evaporation and accurate weighing. Ether extractions
are carried  out in a  hood.
a. Procedure for General Organic Analysis14

(1) Solubility  separation (See Figure 7)

         (a)  Weigh out approximately 0.5 gram of CCE  (a)  in a 50 ml beaker.  As
little as 0.1 g or less can be used; however, the percentage error increases as the weight
of the  aliquot decreases.

         (b)  Add about 1 ml of methanol to the sample  and stir.  Dissolve sample
in 30 ml of ether and stir.  If there is apppreciable insoluble material, filter through
a sintered-glass funnel under vacuum. The residue  (b) is  the  ETHER INSOLUBLE
fraction  (El).  Transfer the residue back to the 50-ml beaker using methanol, evap-
orate on a steam bath, cool and weigh.

         (c)  Transfer the ether  solution  (c) to a 125-ml separatory funnel.  (Do not
use stopcock  lubricants—use only glass  or Teflon  stopcocks).  Extract three times
with 15 ml portions of distilled  water and combine extracts in a tared 125-ml Erlen-
meyer flask.  Evaporate water (d) to dryness on a steam bath with  a jet of air, cool
and weigh. This is the WATER SOLUBLE fraction  (WS).

         (d)  Extract the ether solution  (e) remaining in the funnel three times with
15-ml  portions of dilute HC1 (5%).  Set the ether layer (f) aside  and make the HC1
extract (g) strongly  basic  (pH  > 10) with  NaOH pellets or 25%  NaOH  solution.
After cooling, extract  three times with 15-ml  portions of ether, combine in a 125-ml
14 It is recognized that use of strong acids and bases as described in the solubility separation may
cause condensation, hydrolysis or decomposition reactions to occur.  Thus, specific determinations
may require alternate techniques.

                                        12

-------
                                  WEIGHED SAMPLE (a)
                                           II
                                     add ether, filter
               Ether Solution (c)
               extract with H20
                                        Residue (b)
                                     evaporate, weigh

                                            I
                                 ETHER INSOLUBLES(EI)
     Ether Lcyer(e)
    extract with HCI
                                  Water Layer (d)
                                  evaporate, weigh

                               WATER SOLUBLES(WS)
                                               Water Layer (g)
                                             Make basic, extract
                                                 with ether
     Ether Layer (f)
   extract with NaHC03
             \
      Ether Layer (h)
      dry,  evaporate
        and weigh
       I
Water Layer (i)
(contains
omphoterics.etc)
               Water  Layer (j)
                  make acid
               extract with ether
             Ether Layer (I)
             dry,  evaporate
               and weigh
Water Layer (m)
    Discard
            STRONG  ACIDS (SA)
     Ether Layer (k)
   extract with NaOH
                                                  Water Layer (n)
                                                     make acid
                                                  extract with ether
     Ether Layer (q)
     dry, evaporate
       and weigh

           II
     NEUTRALS (N)
              Ether Layer (o)
              dry, evaporate
                and weigh

                     I
             WEAK ACIDS (WA)
      Water Layer (p)
         Discard
              Figure 7.  Flow Scheme for Solubility Separation of CCE
Erlenmeyer  flask, dry, evaporate and weigh.  The residue (h) is the BASIC FRAC-
TION (B).  Discard the water layer  (i).15
'"'The basic water  layer (i) remaining after ether extraction may contain some amphoteric and
some water-soluble  substances.  If  these substances are of  interest,  a special plan for  analysis
should be set up (34).
                                           13

-------
         (e)  Extract the ether layer (f) three times with 15-ml portions  of .NaHCOa
(5%).  Set the ether layer (k) aside and make  the NaHCO3  extract  (j)  strongly
acidic (pH < 2)  by careful  addition of concentrated HC1.  After cooling, shake vig-
orously to release CO2.  Extract three times with  15-ml portions  of ether, combine
in a  125-ml  Erlenmeyer  flask,  dry,  evaporate,  and weigh.  The  residue (1)  is the
STRONG ACID fraction  (SA).  Discard the water  layer (m).
         (f)  Extract the ether solution (k) three  times with 15-ml  portions of  NaOH
(5%)  and once with distilled water. Caution: Emulsions may  form during this step.
Set the ether layer  (q) aside and make the NaOH  extract (n)  strongly acidic  with
concentrated HC1.  After  cooling, extract three  times with 15-ml  portions of ether,
combine in a 125-ml Erlenmeyer flask, dry,  evaporate and weigh. The residue (o) is
the WEAK ACID fraction (WA). Discard the water layer (p).
         (g)  The ether solution  (q) contains the NEUTRAL fraction  (N).  Place in
a 125-ml Erlenmeyer flask, dry, evaporate and weigh.
         Ether solutions are dried by pouring over a two-inch  column of anhydrous
sodium sulfate followed by ether rinses. Alternately one may add sodium sulfate (10
g) to the flask and filter off the sulfate after standing overnight.

(2) Chromatographic separation of the neutral fraction  (See  Figure 8)
         (a)  Pack activated silica gel16 in a Pyrex glass column 20  mm in diameter to
a height of 10 cm.
         (b)  Weigh17 the  neutral sample in a 10-ml beaker and dissolve in a minimum
amount of ether. Add sufficient silica gel to adsorb the sample.  Evaporate the ether
gently.
         (c)  Wet the column with about 20 ml of iso-octane and add the sample when
the last of the 20 ml reaches the surface of the adsorbent. Rinse the beaker with iso-
octane and add the  rinsings to the column. The beaker should be rinsed several times
with iso-octane and each  succeeding eluent when the eluent is  added to the column.
         (d)  Elute the ALIPHATIC fraction (AL)  with 85 ml of iso-octane18 (a) and
collect in a tared 150 ml beaker.  The eluent should be added carefully with a medicine
dropper  so  as to  disturb  the surface of the adsorbent  as little as possible. A slow,
clropwise, elution rate is  desirable.  It may  be necessary to  apply mild pressure to
obtain a satisfactory rate.
         (e)  After the level of the iso-octane has  reached the surface of the adsorbent,
replace the receiving beaker with another tared 150 ml beaker.  Elute the AROMATIC
fraction  (AR) with 85 ml  of benzene (b)  and collect.
         (f)  After the liquid level of the benzene has reached the surface of the ad-
sorbent,  replace the receiving  beaker with another  tared 150 ml beaker. Elute the
OXYGENATED  fraction (OXY)  with  85 ml  of a 1:1 mixture  of  methanol and
chloroform (c), and collect.
1BDavison Code 950-08-08-226 (60 to 200-mesh), Davison Chemical Co., Baltimore 3, Maryland.
17 It is convenient to retain for future reference about 5 nig of the neutral fraction.
18 Limited investigation suggests that hexane may also be used.

                                        14

-------
                                NEUTRALS
                                  I   I  I
                        Adsorb on Silica gel Column
          a                          b                           c
          I                                                      I
     Elute with                  Elute with                  Elute with
     Iso - octane                  Benzene              Chloroform/Methanol
                                                               (I'D
   ALIPHATICS (AL)           AROMATICS (AR)     OXYGENATED COMPOUNDS
                                                               (OXY)
      Figure 8.   Flow Scheme for Chromatographic Separation of Neutrals


         (g)  Carefully evaporate the three fractions on a steam bath with  a jet of
dry clean air, cool, and  weigh.  The beakers  should be removed from heat and air
before the solvent is completely evaporated.

(3) Infrared spectra

         (a)  Chloroform  extracts—infrared spectra are obtained on selected fractions
that have unusual physical characteristics or odors. In analyses directed at chlorinated
hydrocarbons, the IR  spectra of the aromatic fraction is run.  See Appendix .Three and
Figures 9 and 10.

         (b)  Ethanol  extracts—infrared spectra, only, are obtained on the alcohol ex-
tracts. The  percent recovery of the CCE and concentration (jug/1) are  calculated and
recorded. The percent and concentration of the various fractions,  obtained  through
the separation of the CCE are calculated and recorded.

b.  Column  Chromatographic Separation of CCE (Alternate Procedure)

         This procedure may be employed to  separate  the  aromatic  fraction  more
rapidly from the CCE when this fraction is of interest.  Since most chlorinated hydro-
carbon pesticides, including  lindane, DDT,  DDD, DDE,  dieldrin, endrin, aldrin,
heptachlor  and heptachlor epoxide, are found  in the aromatic  fraction, this step is
employed when pesticide  data are needed and other substances  are  of  lesser  interest.
However, laboratory tests indicate that three other pesticides, methoxychlor and methyl
and ethyl parathion (organophosphorus compounds)  occur in  the oxy fraction  as might
be expected from their chemical structure.

         The use of this  alternate procedure  does not preclude the isolation of the

                                       15

-------
  4000 3000
 100
2000
                            7    8     9    10     11     12    13    14    15
                            WAVELENGTH (MICRONS)
SPECTRUM NO
5AMPLF YAKIMA ftTvro
WASHINGTCH


ORIGIN

PIIPITY
PHASF
THIfKNFSS
IFGFNH
1-
7
DATF
OPFRATOR
RFMABKS
C C H
ARCMfcTTC FRACTTCTi


<
>•
?!
m;

i

Figure 9.  Infrared  Spectrum of Aromatic  Fraction of CCE Sample Supporting
                   Chromatographic Identification of DDT
   4000 3000
  100
 2000
                         1500
CM-i
1000   900
                       800
                                                   700
                             7     8    9    10    11    12    13
                             WAVELENGTH (MICRONS)
SPECTRUM NO.
SAMPLE SAVAHNAH RIVER.
SOUT« CAHOLINA


ORIGIN

PURITY
PHASE
THICKNESS
LEGEND
1
9
DATE
OPERATOR
REMARKS
C.C.B.
AKOHATIC FRACTKX


                                                                        £3
                                                                         i
                                                                         o
Figure 10.  Infrared Spectrum of Aromatic Fraction of CCE Sample Supporting
                  Chromatographic Identification of Dieldrin
                                     16

-------
other classes of organics, because the longer, generally applicable solubility separation
can  be employed with the third  (chloroform-methanol) eluate if necessary.
         This chromatographic separation technique for the CCE is identical to that
used in  separation of the neutral fraction.  In this case, however, 0.5 gram of the CCE
 (or  less) is weighed in a 50-ml beaker, dissolved in a minimum quantity of chloroform
and added to just enough silica gel to adsorb the dissolved sample.  The chloroform is
gently evaporated  and the sample adsorbed on the silica gel is  added to the column
as described  in Section A.2.a.(2)  above.  The aliphatic fraction, the aromatic fraction,
and a complex residue,  eluted with chloroform-methanol, are obtained.

c. Thin  Layer  Chromatographic  (TLC) Separation of Pesticides19 From Aromatic
Fraction of CCE  (38).

(1)  Preparation of plates
         Layers of  silica gel  0.25  mm  thick are prepared  on 200  x  200 mm glass
plates.  A thin slurry is prepared of 30 g of silica gel G in about 60 ml of water and
spread  over five  plates with the  aid of a variable  thickness spreading  device.  The
plates are allowed to stand five  minutes, then dried  in  an oven for  60 minutes at
110° C. and  stored in a desiccator for  future use.20

(2)  Preparing  the solvent system
         The developing solvent,  carbon tetrachloride, is  added  to the chamber to a
depth of 10 mm  (approximately 200 ml).  Two filter paper  wicks, one  on each side of
the  chamber, are placed so that one end contacts the solvent. After the lid is in place,
the  chamber  is allowed to equilibrate for one hour.21

(3)  Spotting of plates
         Marks are made near the edge of each plate  at distances of 1.5 and 11.5 cm
above the bottom edge to define  the spotting line and the point at which the solvent
front has moved to  10 cm.
         The entire  aromatic fraction of the  CCE is dissolved in benzene in a 15-ml
centrifuge tube and  made up  to 0.5 ml.  A 50-ul aliquot or one-tenth of the aromatic
fraction in benzene is spotted22 (see Figure 11).
1!l Samples of very  high total  organic content  (CCE  or  grab) may require  additional clean-up,
e.g., the Mills florisil column procedure (39) (40).
-" Layers of varying thickness have been investigated.-  Layers  thicker than 0.25  mm produced no
better separation and thinner  layers  were consistently less uniform.  Aluminum  oxide plates 0.25-
and 0.50-mm thick  were also prepared.  The  range of Rr values for the  pesticides spotted was con-
siderably smaller than those observed with silica gel plates using the same solvent system. Thus
separation was not as good.  However, alumina plates do have special applications.  For example:
pesticides that show a tendency to streak on silica gel (chlordane, toxaphene)  produce single spots
on alumina.  Also, it is  possible to separate dieldrin and endrin, which  on silica gel normally have
approximately equal Rt  values.
-1 Many different developing systems, both multi-component and single component, have been in-
vestigated. In general,  the  single  component systems showed  much more  consistent results than
did  the  multi-component systems.  The  single component system which  showed the best separation
of all pesticides  investigated was carbon tetrachloride. The range of Rr  values  of pesticides was the
best of all systems investigated.
22 The aromatic  fraction of  the CCE  is  generally 5 to 15 mg, or approximately 1 to 5% of the CCE.
The average  pesticide content  of most river waters  sampled by the  carbon adsorption method has
been estimated to be about 2% of the aromatic fraction. Fifty microliters or 10% of the aromatic
fraction  yields enough  material for good pesticide  peaks in  gas chromatographic analysis, when
separated by  thin layer chromatography.

                                          17

-------
         It is necessary to direct a very  gentle stream of clean, dry air on the point of
spotting to keep the diameter of the spot less than 1.0 cm.  The use of air should  be
kept to a minimum. A typical plate for pesticide analysis  may contain up to  nine
sample spots.  The  samples may  be  spotted  in duplicate;  one for elution and gas
chromatographic  analysis and one for possible corroborative visual identification  on
the sprayed portion  of the plate.  In addition, pesticide standards are spotted  on the
plate.

141 Development

         The spotted plate is placed in the chamber so that the bottom edge is in  con-
tact with the solvent and the lid is replaced. When the solvent reaches the upper ref-
erence line (10  cm),  the plate is  removed and  the solvent allowed to evaporate.  The
spots  are made visible by spraying the developed plate with  a dye or a chromogenic
agent.  Two spraying procedures are described  below.

         (aI RHODAMINE B METHOD

The areas of the plate  containing the unknown samples are masked with a glass plate
and the center area containing the  pesticide standards is sprayed with a fairly heavy
coat of Rhodamine B.  The plate is allowed to dry completely (about 5  min.)  and is
examined under UV  light. The pesticides are seen in natural  light as purple spots on
the pink background,  but are seen much more readily under UV  light where they
appear  as quenched  areas on the  fluorescent background.
                       Figure 11.  Spotting of TLC Plate

                                       IS

-------
        10.0 cm
         8.0 cm
         6.0 cm
         3.8cm
         1.0 cm

SPOTTING 0-7
      LINE  g f
                     ZONE FOR;
                     SPOT I SPOT 2 SPOT 3
               SECTION
                  TZ.
               SECTION
                 33C
                SECTION
                 nr
                SECTION
                 n
SECTION
  I
                           •SOLVENT LINE
                              7.8 ALDR1N
                              7.5 DDE
                              7.3 DDT
                                                 HEPTACHLOR
                            — 5.5 y-CHLORDANE
                            --5.2 ODD
                            --4.2 LINDANE
                                	3.4 HEPTACHLOR  EXPOXIDE
                	2.4ENDRIN, DIELDRIN
 I
I
 SPOT
10   II   12
Figure 12.  Diagram  of Designation of Sections in the Cleanup and Separation
                    of CCE-Aromatics on Silica Gel Layers
        The vertical zones of travel for pesticides present in the unknown samples
will be respectively the same as those of the sprayed standards which are visible. From
this information, the vertical zone of travel for each sample spot is divided into five
horizontal  sections. The sections  are identified with Roman  numerals  as shown in
                                     19

-------
                   A           B         C          D
10
3

m



















*n
\ L












































































|





J
                            Figure 13.  Photograph of a  Developed Thin Layer Plate. A. DDT,  B. Endrin,

                              C.  Heptachlor,  D.  Benzene  Hexachloride,  E. Aldrin,  F.  Dieldrin, G.  ODD,

                              H.  DDE, I. Mixed Standard.

-------
Figure 12.23 A photograph of a developed plate is  shown in Figure 13.  These hori-
zontal sections are determined by the Rf values of the standard pesticide spots.  The
silica gel in each  section is scraped loose from the plate with a spatula.  With the aid
of vacuum, the silica gel, first from the periphery of the spot and then from the area
of the spot itself, is drawn into an eye dropper which is plugged at the tip with glass
wool (Figure 14). The material adsorbed on the silica gel in each eye dropper is eluted
quantitatively  into a graduated 15 ml centrifuge tube with 5 to 10 ml of acetone and
subjected to gas chromatographic analyses.
        The following pesticides have been investigated and can be seen at the 10 /*g
level under UV light after spraying with Rhodamine B:

        endrin                          toxaphene
        dieldrin                         DDE
        DDT                           chlordane
        aldrin                          parathion
        lindane                         methyl parathion
        heptachlor                      ovex
        heptachlor epoxide               tedion
        methoxychlor                    DDD

The Rf values for most of these compounds are listed in Table 1.
      The application of Rhodamine B as a spray reagent has definite advantages
over silver nitrate spray.  With Rhodamine B, the  exact position of the pesticide on
the plate  can be determined without destroying  the pesticide.  Therefore,  using a
selective solvent (ether/petroleum ether 1:1), the pesticide can be eluted from the silica
gel while the Rhodamine B  is retained. However, these advantages apply only when
the entire plate, samples as  well as standards, is  sprayed.
23 Repetitive testing of standard pesticides and the subsequent resolution of nine chlorinated hydro-
carbon pesticides resulted in the designation of the illustrated sections.
The eluate from Section V  was repeatedly analyzed by gas chromatography to determine if chlori-
nated pesticides occurred in it. None of the pesticides studied occurs in this Section. Hence, it is
not analyzed on a routine basis.
                 AIR  FLOW                          A—EYE DROPPER
               •*•
   VACUUM

                          HOSE             ^— GLASS  WOOL
           SILICA  GEL COLLECTION   ASSEMBLY
                   Figure 14.  Silica Gel Collection Assembly

                                       21

-------
         (b) SILVER NITRATE METHOD

An alternate spraying technique (41)  employing silver nitrate may be used for locating
the pesticides on thin layer plates. The samples may be spotted in the manner described
above. However, silver nitrate destroys the pesticide and, thus, if gas chromatographic
analysis  is to follow and corroborative identification on the plate is desired, the samples
must be  spotted in duplicate.  To accomplish this,  two series of  samples are spotted
along with standards. An internal dye,24 previously standardized against pesticides, is
also spotted to determine the zones of travel. The zones are marked and the sections
for the samples to be analyzed by gas chromatography are scraped and eluted as above.
         The samples and standards  remaining  on the plate are then  sprayed  with
silver nitrate, dried, and exposed to UV light until spots appear. Chlorinated pesticides
appear as brown to black spots.
         Chlorinated  and  non-chlorinated pesticides  may be detected by exposing the
untreated, dry plate to bromine vapor for 30 seconds, drying for 30 seconds, and spray-
ing with a  fluorescein solution and finally with silver nitrate (42). Exposure to UV
light for  4 to 7 minutes causes chlorinated pesticides to appear as brown to black spots
and other pesticides  as yellow to white spots on the tan background.
B.   DISCRETE  BOTTLED  SAMPLES

1.  Extraction of Pesticides

a. Extraction From Water

(1)  Semi-Automatie Extraction  (43)  (44)

         Equipment for semi-automatic  liquid-liquid extraction of chlorinated hydro-
carbon pesticides as well as other organic compounds from water  was developed to be
compatible with the sample  bottle currently employed in this laboratory  (see Figures
15 and 16). It consists essentially of two parts. The first part is a cylindrical impeller
housing and bar magnet having  an inlet port and four outlet ports. The second part
is a  plug which fits the  mouth of  the sample bottle and provides the means of recla-
mation of the solvent  when the extraction is  completed.
         Extraction is  accomplished by placing the  impeller in the sample bottle con-
taining approximately 850 ml water, and adding 50 ml of  a hexane-benzene mixture
(9:1). The sample  bottle is capped and inverted on  a magnetic stirrer so that the
impeller  may  operate  on  a flat surface. To reduce magnetic attraction  between the
bottle cap and the bar magnet in the impeller, the center is cut out of the bottle cap.
An aluminum insert with a Teflon liner is used to seal the  opening.
         When the stirrer  is operating, the rotating motion  of the impeller creates a
vertical  vortex  in  the   sample  and  draws  the  hexane-benzene mixture  into  the
central inlet  port.  Small bubbles of the solvent  are ejected  from the four radially
located outlet ports. The bubbles rise through  the outer portion of the sample to the
surface where they collect and recirculate as long as the magnetic stirrer  is operating.
When the extraction has  been  carried out for 30  minutes, the bottle is  removed from
the stirrer. The cap is replaced by the plug used in reclamation  of the solvent.  The
-4 The Rr of the pesticides will vary, since temperature and humidity conditions are not controlled.
However, with the aid of the dye mixture (see Appendix Three) which has been previously stand-
ardized against pesticide spots, the sections may be adjusted to compensate for these deviations.

                                        22

-------
plug is held in place by a pressure device and air under pressure is introduced through
an  inlet.  This air forces  the  hexane-benzene  mixture out of a  tube  which extends
into the solvent phase just above the  solvent-water interface. As  much as possible  of
the solvent is collected and measured  in a 50-ml  graduated cylinder.25
        The water remaining  in the bottle is poured into a 1000-ml graduated cylinder
and its volume measured and recorded.

121 Separately Funnel Extraction
-'•' Recoveries of the pesticides using the semi-automat 11- extractor range from 77 to 95%.
              Figure  15.  Semi-automatic Liquid-Liquid Extractor
                                       23

-------
                     Figure 16.  Solvent Recovery Apparatus


         The entire measured water sample (approximately 1  liter I is drained into a
2-liter separatory funnel equipped with a Teflon stopcock and is extracted successively
with  100, 50, 50, 50, and 50 ml of redistilled hexane.28
         The drained sample container is rinsed with three 50-ml aliquots  of  hexane.
The first two rinse volumes serve as the first extraction  volume (100 mil for the  sample.
The third rinse serves as the second (50 ml) extraction volume.
         The sample is shaken moderately for four minutes.  (Vigorous shaking may
cause severe emulsions,  particularly in waters  of high organic content  and/or high
turbidity. I  The  extracts are combined  in a 300-inl  Erlenmeyer flask and dried  by
-"Other solvents surh as carbon tetrachloride, chloroform, and  ethyl  ether-petroleum  ether (1:1)
may be used  (45) (46) (47).

                                        24

-------
pouring through a two-inch column of anhydrous sodium sulfate. The column is rinsed
three  times with approximately 5 ml of hexane  and the rinsings  are added  to  the
extract.27

b. Extraction From Bottom Samples

(1) Drying the Sample
        The excess water  is decanted and the sample is spread in a pyrex dish  (8"
wide x'12" long x 2" deep). The sample is air dried at room temperature for about 4
to 5 days.  Many pesticides are volatile and  may be lost if drying is  carried out at
elevated temperatures or for an excessive length of time.
        The dried  sample is ground with a  porcelain mortar and pestle to a uniform
particle size.

(2) Extraction
        The sample is divided by mixing and quartering until a sub-sample of about
100 grams  is obtained.  The sample is weighed in a 100-ml  beaker.
        The extraction is  carried'out in  a soxhlet extractor (see  Figure  17).  Glass
wool  (about  1  inch deep)  is packed in the bottom of the  extraction chamber  (40 x
150 mm). The weighed sample is added and  an additional wad of glass wool is placed
on the top.  The sample is then extracted using 200 ml of hexane-acetone  (9:1)  for
about  8 hours.  The  extraction may be  carried out overnight or longer  as may be
necessary for heavily  contaminated samples.
        Several alternate extraction methods are described  in the literature (25) (49)
(50).


2.  Concentration  of Extract

a. Water Sample

        The extract  (approximately 50 ml)  obtained by semi-automatic extraction is
transferred to a 100-ml beaker and evaporated on a 40°C warm water bath to approxi-
mately 4 ml  using a very gentle stream of clean,  dry air. It is then transferred with
rinsing to a 15-ml graduated centrifuge tube  and  evaporated to an appropriate volume
for spotting on a TLC plate.
        The extract (approximately 300 mil  obtained by separatory funnel extraction
is transferred to a  Kuderna-Danish evaporator and concentrated to approximately 4
ml. It is then  transferred  to a 15-ml graduated  centrifuge  tube and evaporated to an
appropriate volume for spotting on a TLC plate.

b. Bottom Sample

        The extract  (approximately 200 ml) is evaporated in the extraction flask
on a 40°C water bath with a jet of clean, dry air  to about 30 ml.  It is then transferred
-" Reported values for efficiency of extraction under these conditions range from 85 to 90% for the
pesticides  (45) (40) (.'tS). It is recommended that each analyst repeatedly check  on extraction
efficiency.

                                       25

-------
Figure 17.  Soxhlet Extraction of Bottom Samples
                     26

-------
to a tared 50-ml beaker  and evaporated to apparent dryness at room temperature.
When possible, the weight of the extract is obtained.
        The extracts are cleaned up  by column chromatography using the method
described  in section A.2.b., page 15. The aromatic fraction is evaporated to 5 to  10
ml and transferred to a 15-ml graduated centrifuge tube and evaporated to an appro-
priate volume for spotting on the TLC plate.

3.  Thin  Layer Chromatography

        The concentrated extract is spotted on a TLC plate and developed in the same
manner as the CCE  aromatic fractions.  The entire extract is spotted whenever pos-
sible. However, care must be taken so that the plate is not overloaded.  Thus, it may
be possible to spot only an  aliquot of  some samples. Samples high in  total organics
are in this category.
        The four eluted sections  are subjected to gas  chromatographic  analysis.
Recoveries of standard pesticides  from the thin layer  ranging from 85 to  98% are
obtainable.28
-8 It is recommended that each analyst repeatedly check the TLC recovery efficiency.
                                       27

-------
IV.   DETERMINATIVE  STEPS
         The identification and measurement of chlorinated hydrocarbons in surface
water require extremely sensitive techniques.  In small samples the low concentrations
which require identification and measurement often provide  only  a few  nanograms
(10~9 gram) of a pesticide. Carbon adsorption samples usually contain larger amounts
for analysis.  Gas  chromatography, thin layer chromatography,  and infrared spec-
troscopy are employed  as corroborative and determinative steps.
A.   GAS  CHROMATOGRAPHY

         Electron capture gas chromatography  (ECGC)  (51)  is used for identification
and measurement because of its sensitivity. Microcoulometric titration gas chromatog-
raphy (MCTGC)  (52), although less sensitive, is specific for halogenated substances.
The  use of both systems combines the advantages  of  specificity and  sensitivity.29
Sample gas chromatographic traces for the ECGC and MCTGC systems are included
in Appendix Two.


1.  Application  of Electron Capture Gas Chromatography

a. Extracts of TLC  Sections of CCE Aromatics
         Pesticides from  each sample  separated by  thin layer chromatography  are
contained in acetone in "four  15-ml, calibrated, Pyrex centrifuge tubes.  The volume,
usually three to five ml, is reduced by evaporation to 0.5 ml in a water bath at 40°C.
with a jet of clean  dry air.  A 5-/J.1 aliquot is withdrawn  from each tube  with a 10-^1
Hamilton microsyringe and injected into the previously conditioned and  stabilized
column.30  Although  these  conditions  are adequate  for the  concentration range of
pesticides found  in most  samples, in some instances  the volume of the  TLC extract
must  be adjusted by evaporation in a 40° C. water bath or by dilution. It is often
possible to predict  the need for  adjustment of injection volume or the total  volume
on the basis of the  TLC result.
-9 Some other detectors used for determination of chlorinated and/or thiophosphate pesticides are
the  sodium  thermionic (53)  (54),  stacked  flame (55), flame photometric (56), emission spectro-
photometrie (57), and the Burchfield microcoulometric titration cell (58).
•"•"The instruments employed are Perkin-Elmer models 154—L and 811 equipped with  a parallel
plate E.G.  detector, pulser, D.C. power supply, amplifier, and Leeds &  Northrup (0-5 mv) re-
corder.  They are operated with pyrex glass or aluminum columns, 4 ft. long x %-inch O.D., packed
with 5% D.C. 200 silicone on 60/80 mesh acid-washed  Chromosorb P  at temperatures  of 180 to
195°C. The power supply is operated in the pulse  mode and  the carrier gas is 95%  argon—5%
methane at a flow rate of 120 ml/min.  (nitrogen carrier gas is used for the D.C. mode). The am-
plifier is usually operated at an attenuation range of 1 and an attenuation of 16 to 64.

                                        28

-------
        After the chromatogram is obtained, it is examined for peaks which possess
retention times and peak geometries which  match known pesticides  (see Appendix
Two, Table 2).  Since the aromatic fraction has been separated by TLC, the number
of possible pesticides in a given injection is reduced. The areas under these peaks are
calculated and compared to a standard calibration curve  which is prepared by  ob-
taining peak areas from known quantities of the individual pesticides under identical
conditions. The peak areas are measured as illustrated in  Figure  18  or  by use of  a
planimeter, disc integrator, or electronic digital integrator.
        The calibration curve is obtained  by  plotting peak  area in square inches
against  sample  size in  nanograms (see Appendix Two).  As seen in  the discus-
sion of  thin layer  chromatography, three  curves are  necessary  for TLC  II,  three
curves  for TLC III, and four curves for TLC IV.  From  the  calibration curve,  the
nanograms of each pesticide per TLC section, W, is calculated.
            TLC extract)  (ng determined/injection)
                      (/*! injected)
= ng/TLC section = W
b.  Extracts of TLC Sections from Solvent Extract of Bottled Samples

        Essentially the same  procedure as outlined in IV.A.la is  employed except
for the changes noted  herein.  The TLC extracts of sections II, III, and IV are re-
duced in volume to 0.2 ml. A 5-/A aliquot is  chromatographed with an  attenuation
of less than 64.
                       B
                                 AREA UNDER  PEAK -  a  x C

                                 WHERE'

                                    c  = WIDTH  AT  1/2 HEIGHT

                                    a  -= HEIGHT OF PEAK
                       A    D
                CALCULATION  OF  PEAK  AREA
                    Figure 18.  Calculation of Peak Area
                                     29

-------
         The nanograms of each pesticide determined  per TLC  section, W, is  calcu-
lated.

         (til TLC extract)  (ng determined/injection)       /mT „    ,.      ,TT
                                         	*	  = ng/1LC section = W
                         (/tl injected)


2.  Application of Microcoulometric  Titration Gas  Chromatography

a.  Extracts of TLC Sections of CCE-Aromatics

         In many instances the entire remainder of  the extracts of sections II,  III
and IV must be injected in order to elicit a response within the sensitivity of the  in-
strument.  The result of the ECGC run is used as a guide in the choice of an injection
volume. The volume selected is injected into the microcoulometric titration gas  chro-
matographic system.31   The procedure  for identifying  and measuring pesticides from
the gas chromatographic traces  are identical to  those described for the ECGC system
(see Appendix Two, Table 2).   The nanograms per TLC section, W, are calculated
as previously.

         (/u,l TLC extract)  (ng determined/injection) 	    /TIP    f   	W
                         (fi\ injected)                     ^      S6C

b.  Extracts of TLC Sections From Solvent Extracts of Bottled Samples

         Using the result of the  ECGC injection  as a guide,  an aliquot of  extracts
II, III, and IV is selected and injected under the  same  conditions  as  described  for
the CCE-aromatic-TLC extract  in  IV.A.2.a. The evaluation of the chromatographic
trace in terms of identification and measurement  is carried out as described previously.

       (p.1 TLC  extract)  (ng determined/injection)  	    'TT C   t'   	W
                       (/J injected)


3.  Calculations

a.  Bottled Samples


                     •(vlMV.MvlT'cE) =^Pesticide/liter
   Vwx = volume of solvent representing extract of water sample               (/xl)
   Vsp  = volume of Vwx which was spotted on  TLC                           (/J)
   Vt   = volume of solvent representing extract of TLC section               (/J)
   V|   = volume of Vt which was injected                                    (/tl)
   Vsa  = volume of water from which extract was made                       (ml)
   Wj  = weight of pesticide determined in  injection, Vi                       (ng)
   E    = efficiency of extraction                                         (decimal)
"i The instruments employed are Micro-Tek models 2503R and 179 DSS equipped with a S-100
furnace, T-300 titration cell, and a C-200 coulometer (Dohrmann Inst., Mountain View, California),
and a Minneapolis-Honeywell Brown (0-1 mv) recorder equipped with a disc integrator. They are op-
erated with pyrex glass or aluminum columns, 4 ft. long x %-inch O.D.,  packed with 5% DC 200
silicone on 60/80 mesh acid-washed chromosorb P, at a temperature of 200°C, with a carrier gas of
helium at a flow rate  of  120 ml/min. The injection block temperature is  240°C.  The coulometer is
usually operated at a sensitivity of  200 to 300 ohms.

                                         30

-------
         (1)  If the entire volume of the orginal extract, Vwx is spotted on TLC, then
Vw
•^•*  equals 1 and the term is eliminated.
Vsp
Then,           '^ Mg Pesticide/liter
If, in addition, the weight (/xg) of pesticide/TLC section,  W,  has been  determined
previously :
                                 w_    t   (W.)
                                           V,
Then the calculation reduces to:
                             (W)
                           (V8a)  (E)
                                     = jug pesticide/liter
b.  Carbon Adsorption Samples .
                 (V.t) (Vt) (W.) (C.t)  (10-°)  _;AgDesticide/liter32
                       (V.p)  (V,) (Waf)       — rt5 pesticide, liter

   Wa( ==  weight of aromatic fraction                                         (mg)
   Ca(  =  concentration of aromatic fraction                                (/*g/l)
   Vaf  =  volume of dissolved aromatic fraction                                (/xl)
   Vsp  ==  volume of V»f which was spotted on TLC                             (/u.1)
   Vt   =  volume of solvent representing extract of TLC Section                (p.1)
   Vi   =  volume of Vt which was injected                                     (/*!)
   Wt  =  weight of pesticide determined in injection, Vi                      (ng)

If the weight (ng) of the  pesticide/TLC Section, W, has been determined previously,

                                  w_(Vt)
                                     ~    (V,)
then the calculation reduces to:

                    (W)  (V.f) (Caf) (1
                               (Wat)
                                          = /*g pesticide/liter32
4.  Column  Packings

         Glass and aluminum gas chromatographic columns are used for routine anal-
ysis of chlorinated pesticides.  These columns can be  expected to give satisfactory re-
sults up to six months. The types of injections  made  include:  pesticide standards,
eluates  of  TLC sections of CCE aromatic  fractions from raw and finished waters, and
solvent  extracts of grab samples of raw and finished water.  Other column  packings
:i2 In the light of the unknown efficiency of adsorption on and desorption from carbon for most or-
ganic compounds, these concentrations must be considered minimum; the actual concentration be-
ing equal to or most likely greater than that determined.

                                         31

-------
have been employed in our laboratory. Some additional  column packings  used for
pesticide analysis are shown in Appendix Two, Table 3.  It is recommended that sev-
eral different column packings be employed for corroborative qualitative identification.


5.  Column Conditioning

         To obtain  optimum response and peak  resolution, gas chromatographic col-
umns must be  adequately  conditioned.  Conditioning requirements vary for  different
column packings and for different pesticides.
         Columns packed with acid washed  Chromosorb P (60-80 mesh)  coated with
5% DC 200 silicone are conditioned, in our laboratory, according to the following pro-
cedure. The column is  installed in the oven, the  carrier gas is adjusted to the proper
flow rate, and  the column temperature, increased very slowly to 200°C. (programmed
at a rate of 1°C. per min.).
         It is held at 200°C. overnight and then increased to 225°C. for two to  three
hours and brought back to 200°C.  The column is held at this temperature and 50 to
100  //.g quantities of standard  pesticides and normal  aliquots of actual samples  are
periodically injected.  Conditioning  for the  electron capture gas chromatograph may
require up to three days. Approximately seven days may be  required for the micro-
coulometric gas chromatograph.  For some pesticides, such as  endrin, the column may
require additional conditioning.
         Columns tend  to lose their  response and  resolution abruptly and new columns
should be pre-conditioned and ready for use when this occurs.  Insertion of quartz wool
ahead of the column will tend to lengthen the life of the column by retaining  the non-
volatile waxes  and oils  present in  many of the  samples.  Insertion of a Pyrex glass or
quartz tube in  the injection block is also helpful in extending column life.
B.  INFRARED  SPECTROPHOTOMETRY

    The aromatic fraction of each carbon adsorption sample is prepared for infrared
spectroscopy (13).  A portion of the aromatic fraction is diluted with  chloroform.  A
suitable volume is spread evenly on a simple salt  plate.  The solvent  is evaporated
under a heat lamp.  A standard 12-minute scan  is made on  a Perkin-Elmer  137-B
Infracord33.  The resulting spectrum is examined for specific absorption bands coin-
cident with those appearing on standard spectra (see  Appendix  Three).  When pesti-
cides are present in overriding concentrations, spectra are specific enough to permit
unilateral identification.  In  many cases the number and position of  the absorption
peaks in the  spectrum will lend  support to the identifications  made by previously de-
scribed  chromatography  (see Figures 9 and 10).
        When  sufficient  extract is  obtained from a bottled water or bottom sample,
it is also examined by infrared as described above.
°13 More refined I.R. instruments, equipped with a beam condenser and scale expansion, will provide
much greater sensitivity.
                                       32

-------
V.   CONTROL  OF  INTERFERENCES
        When using ultrasensitive analytical techniques, particularly electron capture
gas chromatography for pesticide analyses, possible interferences from solvents,  acti-
vated carbon, and other reagents  and materials employed throughout the procedure
must be given continuous  attention.  Adequate steps  must be taken to eliminate or
minimize any interferences and ensure that,  if present, they  are taken into consider-
ation in the interpretations of the gas chromatograms.
        To accomplish these objectives, all solvents,  carbon  and other materials are
checked routinely by subjecting them to analyses identical to those used for samples.
A.   SOLVENT  INTERFERENCES


1.  Chloroform

        All chloroform (analytical reagent grade) used for extraction of the carbon
adsorption  samples  is distilled before use. To determine interferences, a volume of
CHC18 equivalent to that used for  extraction of the carbon is distilled a second time.
The residue obtained, representing that attributable to CHCls which would have been
contained in the CCE  sample is given  the column chromatographic separation  [see
Section III.A.2a(2)]. One-tenth of the  aromatic fraction thus  obtained  is given  the
usual TLC cleanup. A  volume of the eluate from each section,  equivalent to  the vol-
ume of sample routinely used, is injected into both  the electron capture and the micro-
coulometric  gas chromatographs. The significance of interferences, if present,  is noted
in terms  of retention time, peak geometry and peak intensity, i.e., area  and height.
Interferences, if noted under these conditions, can be considered maximum.


2.  Hexane-Benzene

        Chromatographic grade hexane and benzene are checked individually as below
and, if necessary, distilled in an all-glass system before use in extraction of water grab
samples.  A  volume  of  hexane-benzene  (50 ml, 9:1) or hexane (300 ml)  equivalent.to
that used in the  extraction is evaporated to the appropriate  volume as  described in
Section III,B,2. This residue is given the TLC cleanup and the  eluates from the four
sections are injected into the gas chromatograph as discussed previously.
        Other solvents or solvent combinations (ethyl ether, petroleum ether, benzene,
acetone, chloroform, and carbon tetrachloride) used for extraction  are checked in  the
same manner.

                                      33

-------
3.  Hexane-Acetone

         A volume of hexane-acetone (200 ml, 9:1) equivalent to that used in the ex-
traction  of a bottom sample is evaporated as described in section III, B,l,b. The resi-
due is examined by TLC and gas chromatography.


4.  Carbon Tetrachloride and Acetone

         A quantity  of carbon tetrachloride and acetone approximately equivalent to
that used for development of the TLC plates and elution  of the  pesticides from the
silica gel are also checked for interferences.
B.   CARBON INTERFERENCES

         As discussed in Section II entitled Sample Collection, it is possible for small
amounts of organic substances to become adsorbed upon carbon in the period between
its activation and  its use in the cartridge.  Together with a precautionary program to
reduce the probability of such contamination occurring during transport or storage, an
aliquot of the unused carbon  is analyzed by a procedure identical to that used for car-
bon adsorption samples.


1.  Carbon Blank

         The presence  or absence of interferences in the carbon blank are determined
according to the procedure for determining the CCE  (Section III.A.l.). A quantity of
carbon equivalent  to that  used in a carbon adsorption cartridge is extracted with three
liters of  double distilled analytical reagent grade chloroform. A blank determination
is made  whenever  a new container is opened.  All  cartridges filled with carbon from
a given container are so recorded.  The residue  is subjected to the standard  column
chromatographic separation.  One-tenth of the  aromatic fraction is given the normal
TLC  cleanup and  gas  chromatographic  analysis as previously described.
         Interferences,  if noted under these conditions, would be at maximum effect.
C.   OTHER  SOURCES  OF  INTERFERENCE

        The silica gel (Davison Code 950)  used for the column chromatographic sep-
aration, silica gel G used for TLC cleanup, and the anhydrous sodium sulfate used for
drying solvent extracts are examined  for interferences  using the appropriate solvents
for  elution and  development.  The  eluates are subjected to gas chromatography as
previously described.  When quantities of residue permit, infrared  spectra are deter-
mined  for solvents, carbon, and other reagents.
        The following techniques  are suggested for pretreatment and clean-up of the
adsorbents used  for column and thin layer chromatography. Silica gel (Davison) may

                                       34

-------
be washed with acetone-ether (1:1) followed by  ether, then air dried,  and finally
heated at 110-120°C. overnight (59).  Pretreatment  of  florisil may  include  washing
with acid, alkali, absolute methanol, and distilled water  followed by heating at 650°C.
for 1-3 hours.  Spent florisil may be recovered by  washing with diethyl ether,  benzene,
ethanol, and distilled water then  heating overnight at 130°C. (60).  Silica gel G arid
alumina TLC  plates may be washed by developing with distilled water and drying at
75°C. for  15 minutes. The washing is repeated and the  plate is dried at  75°C.  for 30
minutes (61).  Alternately, 50% aqueous acetone may be  used for washing.  The plate is
air dried for 5 minutes and then heated at 80°C for 45 minutes (62).  The adsorbents
prepared by these techniques may be stored in a desiccator from 4 to 7 days without
significant loss of  activity.
        Glassware may  also be a source of contamination. Therefore all glassware is
cleaned up as  described in Section II.B.3. If the type and size of the glassware  per-
mits, it is  heated in a muffle furnace at about 400°C. for 15 to 30 minutes.
D.   INTERPRETATION

           The interpretation of all gas  chromatographic analyses are made in light
of any interferences determined by the foregoing procedures.  If interferences are pres-
ent and are significant  enough to invalidate  specific 'results,  either qualitatively or
quantitatively, these results  are discarded.
                                       35

-------
VI.   SENSITIVITY  AND  SPECIFICITY
A.  SENSITIVITY

         In discussing sensitivity, in terms  of concentration, it must be  pointed out
that concentrations  obtained  with the  carbon adsorption method are minimum values
(see Section IV).


1.  Carbon Adsorption Extracts Examined by Electron Capture Gas Chromatog-
    raphy

         The electron capture detector varies in its sensitivity for the various members
of the chlorinated hydrocarbon pesticide series. However, it is, in  general, capable  of
detecting absolute quantities of 1 nanogram  (1CP9 g) or less on a routine basis. Remem-
bering that (a)  only 1/10 of the aromatic fraction is subjected to TLC and (b) that
only 1/1000" of the TLC section extract is chromatographed, then  under these  condi-
tions a peak calculated to represent 1 nanogram is equivalent to:

                       1 ng X 10  X 100 = 1,000 ng =  1 /xg

         If a sample had a measured volume of 5,000 gallons  (approximately  20,000
liters) the lowest detectable concentration  can be estimated at 1 jug/20,000 liters  or
0.00005 ju.g/1. However, if it is also recalled  that (c)  only a portion  (variable)  of the
CCE is separated by solubility to obtain the neutral fraction and (d) that only a por-
tion of the neutral fraction  is separated on the silica gel column to obtain the aromatic
fraction, the lowest  detectable concentration is greater than 0.00005 n*g/l.  Since the
values for  (c) and (d)  vary, the lowest detectable concentration,  under these  condi-
tions, has been  estimated conservatively  at 0.001 /*g/l.  Samples measuring less than
5,000 gallons induce a  corresponding increase in the estimate of the lowest detectable
concentration.  It  must be noted, however, that the electron capture detector sensitiv-
ity can often be used to detect quantities  of less than 1 nanogram  for some pesticides
if  background interference is negligible.
         In the  event  (a) it is necessary  to detect concentrations of less than 0.001
jug/1, (b) the additional effort is justified, and (c) interferences are negligible, the use
of all of the CCE, all of the neutral fraction, all of the aromatic fraction, and all of the
TLC section extract has a potential for detecting a concentration  of 0.00000005 /u.g/1
in a 5,000-gallon sample and 0.00000025 jug/1 in a 1,000-gallon sample.

                                        36

-------
2.  Carbon Adsorption  Extract Examined by Microcoulometric Titration  Gas
    Chromatography

        Since 2-10  ng are required  to produce a minimum recognizable response for
most chlorinated hydrocarbon  pesticides and recalling that it is usually necessary to
inject all of the TLC section extract (rather than 1/100 as in the case of ECGC) the
lowest  detectable concentration under these  procedures has been conservatively esti-
mated  at 0.001 /*g/l.  Potentially, using the  entire sample  and with significant addi-
tional effort, detection of 0.0000025 /xg/1 is possible.


3.  Bottled Sample Extracts Examined by Electron Capture Gas  Chromatography

        The lowest measurable concentration is approximately 0.001 ftg/1  in a 1-liter
water sample and 0.001 /ug/100 g or 0.010 /*g/Kg in  a bottom sample.


4.  Bottled Sample Extracts Examined by  Microcoulometric Titration Gas Chro-
    matography

        The lowest measurable concentration is 0.002-0.010 /u,g/l  in a  1-liter water
sample and 0.002-0.010 /tg/100 g or  0.02-0.10 /*g/Kg in a bottom sample.
B.  SPECIFICITY


1.  Carbon Adsorption Samples

        In the examination of CCE for chlorinated hydrocarbon  pesticides  by the
procedure outlined, it is demonstrated  that the pesticide:  1—is adsorbed  on  carbon,
2—is  desorbed  with chloroform, 3—is  ether soluble, 4—is not water  soluble,  5—is
not acidic, 6—is not basic, 7—is neutral, 8—is benzene soluble, 9—moves on TLC  in
the same fashion as a given standard, 10—is eluted from ECGC at the same retention
time as, and having the same peak geometry as  a  given standard, 11—is identical  to
the same standard when  chromatographed with  MCTGC in terms of its retention
time,  peak geometry, and degree of chlorination, and produces an infrared spectrum
which in many cases supports the identifications made by  Chromatography.


2.  Bottled Samples

        The  examination of bottled samples by the procedure outlined provides for
three  corroborative chromatographic identifications  which  serve as a three-way cross
check on identification.
                                      37

-------
                         APPENDIX  ONE
              ENGINEERING ASPECTS OF  SAMPLING
             BY THE CARBON ADSORPTION  METHOD
        The carbon  adsorption method of organics sampling consists of the passage
of up to 5,000 gallons of raw water at rates up to % gallon per minute through a carbon
adsorption column. Following the sample run, the column is shipped to the laboratories
in Cincinnati,  Ohio, for analysis.
I.  TYPES OF SAMPLING  EQUIPMENT  IN USE
A.  GENERAL

        At the present time there are three types of carbon adsorption sampling ap-
paratus  used  in the  Water Pollution  Surveillance System. The first and oldest of
these consists  of a piping  arrangement that was originally assembled and installed
at the sampling location. This device is referred to as  the manual type installation
and is discussed on page 41. The second type is  a prefabricated system with auto-
matic backwash of a sand prefilter. Two models of this type  system were developed
for extensive use  in water  pollution surveillance. One model is a panel unit equipped
with automatic backwash  device, designed for mounting on a wall inside a building.

                                   38

-------
        Figure 19.  Carbon Adsorption Column and Shipping  Container
A second  model is similar to the panel unit, hut is built into a protective housing for
operation  in remote or outside the plant use.  The  third and  newest  type of  sampler
utilizes a  low flow rate for more efficient collection of organic substances.  Two models
of this type of sampling system  have been designed and  recently placed in  use fol-
lowing successful field evaluation.
B.  DESCRIPTION  OF CARBON  ADSORPTION  COLUMN  (CAC)

        The CAC consists of a piece of Pyrex glass pipe 3 inches in diameter and 18
indies  long.  The ends arc-  fitted  with  brass  plates and %-inch galvanized  nipples.
A  stainless steel screen is fixed in  a  neoprene gasket  at both  ends.  The filter unit
arrives at the station packed with  activated carbon  ready for use.  A special shipping
container  is provided for  returning the  filter.  The  unit, with the shipping container,
is  shown  in Figure 19. A modified cartridge  with  a hose  type connection has been
designed  for use with  the low flow rate  sampling equipment.

                                      39

-------
      16-18  MESH
      STAINLESS STEEL
      SCREEN
  STANDARD  3 PIPE
  THREADED  EACH
  END
16-18 MESH
STAINLESS STEEL
SCREEN,SHAPED
TO FIT REDUCER
COUPLING
                                cvj
                                          3" XI" REDUCER COUPLING
                                    PACKED  WITH GRAVEL
                                                 --    GRAVEL
                                        0.6 TO 0.8 mm SAND
                                                    -    GRAVEL
                                           3" x |"  REDUCER  COUPLING
                    Figure 20.  Details of Sand Prefilter
C.  PRESETTLING AND  PREFILTERING

        Turbid river waters frequently clog the CAC when attempting to sample 5000
gallons.  To permit this amount of water to pass  through the column a presettling
tank and prefilter containing sand and  gravel were sometimes required for the man-
ual  and  automatic backwash sampling  systems designed for use at 0.25 to 0.50 gpm
flow rates. This equipment has  not been required for  satisfactory  operation  of the
newer low flow rate designs. A standard hot water tank connected  with the inlet at
the  bottom  and outlet at the top and with a  clean-out tap at the  bottom can serve
as a presettling tank.  The outlet is connected to the  prefilter  containing sand and
gravel. The tank must be flushed at  intervals to prevent accumulation of solids.
        The sand prefilter consists of a steel  pipe  3  feet  long and 3 inches  in di-
ameter, threaded at both  ends,  and equipped  with  3  by 1-inch reducer couplings.
Two cupped stainless  steel screens  are  fitted to the reducer couplings. The space be-
tween the screens is packed with gravel and sand  as shown in Figure 20.
                                   40

-------











.



SAND PREFILTER — *•




f
(IF R
RAW
WATER
PRE-




UM
EQU

SET



C





V
^
9
IRED)

V

<-
— VALVE
f 	 VALVE
X >v






-UNION





J '->-*-iiNinN
1-

TLING^
(IF REQUIRED)
}•






^




PRESSURE
(




GAUGE c

CXI
-•—VALVE \
FLOW \
REGULATOR-3
(1/2 GPM)
•*— VALVE



_
•« 	 VALVE
r- VALVE
y
X A

r- UNION \ 	 WATER
METER
CARBON
•* — ADSORPTION
COLUMN
T"p
\. p-«— UNION
]

t
-
•*— VALVE
/— DRAIN
**>t — DRAIN
    Figure 21.  Schematic Diagram of an Installation with Manual Backwash
D.  INSTALLATIONS WITH  MANUAL BACKWASH
        The presetting tank, the sand prefilter and the CAC are installed at the most
convenient source of raw water. If less than 15 psi pressure is available, it may be
necessary  to pump the water through the system. A drawing of a workable system is
shown  in  Figure 21.
        A water meter located at the end of the system is used to measure the volume
of water sampled.  This  is usually  a disc-type meter, or oscillating piston-type meter,
registering in gallons and capable of measuring  flows as low as ^ gallon per minute.
If necessary, a ^-gpm flow regulator or a valve following the meter can  be  used to
control the flow  rate.
        Fine carbon dust washes out of the CAC when it is first started.  A few gal-
lons of water are passed through the top connection and through  the CAC drain be-
fore the meter is cut  in, to keep the meter  free of the carbon.

                                      41

-------
Figure 22.  Carbon Adsorption  Unit Model  H2O-M1C with Sand Prefilter and
                          Automatic Backwash

-------
Figure 23.  Carbon Adsorption Unit Model H2O-M2C with Presettling Tank and
                        Auxiliary Equipment in Shelter
E.  INSTALLATIONS  WITH  AUTOMATIC  BACKWASH
        Preassembled panel units with automatic backwash of the sand prefilter were
developed to ease installation and operation of the organics sampling apparatus.  Fig-
ure 22 shows the Model HaO-MlC panel unit  designed for installation in water treat-
ment plants and other buildings. This equipment has an electric timer and solenoid
valves to backwash automatically the sand prefilter.  The  panel includes an electric

                                     43

-------
                                   Organics  Sampler
                          Models H20 - MIC and  H20  - M2C
  A) Sample Type - Groanics
  B) Sampling Volume - 5000 Gallons
  C) Sampling  Frequency - Monthly
  D) Information -
                U.S. DEPARTMENT OF INTERIOR
                Federal Water Pollution Control Administration
                Water Quality Activities
                1014 Broadway
                Cincinnati 2, Ohio
         SCHEMATIC FLOW DIAGRAI
             NORMAL
   LEGEND        FLOW BACKWASH
    -SOLENOID VALVE —CLOSED  OPEN
    - SOLENOID VALVE - OPEN  CLOSED
    - SOLENOID VALVE - OPEN  CLOSEO
    - SOLENOID VALVE — CLOSED  OPEN
    -I apn FLOW CONTROL VALVE
    - 2 gpm FLOW CONTROL VALVE
    -PRESSURE RELIEF VALVE
    -MANUAL CONTROL VALVE
    -GRAB SAMPLE VALVE
    -PRESSURE GAUGE
   [>- 3-WAY COCK
   -* NORMAL FLOW
   ~~ BACKWASH
SAMPLING PROCEDURE

I) Install carbon filter
2) Flush carbon filter, in place through cock
  "X", with raw river water to remove
  carbon fines.
3) Begin sampling run and record date and initial
  water meter reading on carbon filter log sheet
*) Record daily, if possible, the water meter
  readings on the carbon filter log sheet until
  the required volume of raw water has been
  sampled. (Approximately 7 days)
5) After filtering required volume record removal
  date, water meter reading and return carbon
  filter and log sheet to Cincinnati, Ohio
  within the provided shipping container
6) Upon receipt of used carbon filter in
  Cincinnati, a new filter will be returned for
  next sampling run
Note:  In highly turbid waters, the filter may
     tend to clog, turning filter end to end
     and/or backwash ing for 2 to 3 minutes may
     be used once to obtain at least a 2000
     gallon sample
Figure  24.  Schematic  Flow  Diagram  with  Sampling  Procedure for  Organic
                     Sampler Models H2O-M1C and H2O-M2C
disconnect switch  of  fuse-plug-type  and  grounding-type duplex outlet for pump.  It
also has three 3-way  cocks,  one to  protect  the water  meter from fine carbon at the
beginning of  sampling, one to facilitate  checking of the flow control valve and water
meter,  and  one to  check  backwash  performance.
         For remote locations the sampling apparatus is installed in an insulated equip-
ment shelter.  An organic sampling field  unit, Model H20-M2C, containing preassem-
bled panel apparatus,  a 30-gallon presettling tank,  electric space heater, and auxil-
iary equipment is shown  in Figure 23.  The  pumping system will vary depending  on
the needs of the individual sampling station.  A submersible pump  was used for the
field unit  shown in  Figure 23. The equipment shelter has space for  a  jet centrifugal-
type pump, or other acceptable motor pump unit.
         A prefabricated metal building may be  provided, where required, to  provide
a permanent shelter for equipment and operating  personnel. This type of building is
usually  installed  on  a reinforced concrete  base.  An organics-sampling  panel  unit
(Model H20-M1C), a pumping  system,  and other  sampling  equipment can  be  in-
stalled  in this type of facility.
         Figure 24 shows the  schematic flow diagram with sampling  procedure for the
organics  sampler.  (Models H20-M1C and H2'0-M2C.)
F.  LOW FLOW  RATE  ORGANICS  SAMPLING
          Studies of optimum sampling  rate and sample volume for maximum recov-

                                           44

-------
ery of organics by the standard carbon adsorption method (9) showed that sampling
efficiency can be increased by the use  of smaller volumes and lower flow rates. Cas-
telli and Booth (30) designed a practical system to control flow  of raw water through
the carbon column at low  rates and measure the throughput.  This system was devel-
oped further by Reid and  Stierli (31) for field evaluation.
1.  Comparative  Field Tests

        A preliminary field evaluation of two low flow rate samplers in comparison
with  conventional  sampling apparatus  was conducted  at the Surveillance  System
Field Test Station  on  the Little Miami  River, Cincinnati, Ohio, during February and
March 1964.  Four  sampling panel units were operated in  parallel, two  conventional
units at the rate of %-gpm  flow rate and two prototype samplers at the reduced rate
of 100 ml/min, or less. Sample volumes  for the higher flow rate were approximately
5000  and  1000 gallons and  about  250 gallons (1000 liters) for the lower rates.  Figure
25 shows  the  apparatus  as  it was installed at the Field Test Station. The panels in
the foreground and left background operated at l/% gpm, while the other two  panels
(one  behind the center panel and the other in the right background) operated at the
reduced rates of flow.
        Included in the study were tests with and without presettling and sand pre-
filtering. The  performance of the low  flow rate equipment was satisfactory without
presettling and prefiltering  even though turbidities of up to 1750 Jcu were measured
for the raw water.
        The study  indicated that approximately double the  amount of total organic
materials  is recovered per  gallon  of  water passing through  the  "regular"34  carbon
column by reducing the  throughput  from 5000 to  1000  gallons.  Approximately five
times the  amount of total organic substances was  recovered  per gallon of water by
decreasing the  rate  flow from y2 gpm to 100  ml/min and reducing the throughput
from  5000 gallons to approximately 1000 liters. The use of "all fines"35 carbon col-
umns further increased the  recovery of  organic materials  when sampling at conven-
tional and low flow rates.
        The comparative tests with and without  presettling and prefiltering indicate
significantly greater recovery of organic  materials from raw water samples receiving
no processing prior to  flow  through the carbon column. Six to eight times the amount
of total organic substances per  gallon of water  were recovered from "all fines"  carbon
columns operated  with low  flow rates and no preprocessing of turbid water as com-
pared  with parallel  "regular" carbon  columns receiving presettled and prefiltered wa-
ter at a flow rate of */2 gpm and a 5000-gallon throughput.   The filtering  action  of
either "all fines"  or "regular" carbon  columns  can be utilized in the low flow rate
.samplers to obtain  an  organics sample which includes much of the silt and other par-
ticulate matter transported by a river. This is  of special importance for measurement
of pesticides in water as the transported  material  may carry specific substances  of
concern.
        Additional field tests of low flow rate samplers were  conducted  on the Mis-
34 The "regular" carbon column is packed with two types of carbon (see page 10.)
:ir> The "all fines" carbon column is packed only with 30 mesh Nuchar C-190 carbon. (See page 10.)

                                       45

-------
souri River at Omaha, Nebraska, the Arkansas River  at Little Rock, Arkansas, the
Columbia  River at  Pasco,  Washington,  and the Escambia River at Century, Florida.
Over 30 production  model low flow rate samplers  were installed during 1965 for wa-
ter pollution surveillance.
2.  Low Flow Rate Organics Sampler, Model LF-1

        Figure  26 shows a Model LF-1 organic* sampler for water. The carbon col-
     and accessory equipment is assembled on a plywood panel 2'6" wide by 3' high.
1111IM
Figure 25.  Equipment  Installed in Field Test Station for Field Evaluation of Low
    Flow Rate Samples in Comparison with Conventional Sampling Apparatus
                                      46

-------
           Figure 26.  Low Flow Rate Organics Sampler, Model LF-1
Raw water enters the sampling system at the left and passes through a 1-gpm flow con-
trol valvejf located behind the pressure gauge. An adjustable pressure relief valve to the
left of the pressure gauge is used to control  the pressure for operation within 3 to 15
psi.  The sample water passes through a Teflon tube  to  the carbon  column.  After
passing up through this column, the water flows through a rubber hose  to a  peristaltic
action type pump  for control of flow at  approximately 100 inl/min.
        The  water goes from the pump to the volumetric measuring tank which con-

                                       17

-------
           Figure 27.  Low Flow Rate Organics Sampler, Model LF-2
tains probes for control of the solenoid valve  below  it.  When water  in  this  tank
reaches  the  top probe it  activates  the  liquid level control and solenoid valve to drain
one liter of water and close the valve.  This volume is automatically recorded in liters
on the digital  counter at the top of the panel. Normally, a one-week sampling period
is used  to collect a sample with approximately  1000  liters throughput.

-------
3.  Low Flow Rate Orgcmics Sampler, Model  LF-2

        A Model LF-2 organics sampler is shown on Figure 27. This sampler is sim-
ilar to the Model LF-1 sampler except it includes a constant head tank between the
carbon column and • peristaltic  action type pump.  A float in the  constant head tank
controls the flow  of water from the carbon  column.  The flow through  the  constant
head tank is regulated by the pump during operation.
        The volumetric  tank,  liquid  level control, solenoid valve, digital counter
and fused electric  disconnect switch operate similarly on Models  LF-1 and LF-2.
        Satisfactory operation  can be obtained with the Model  LF-2 sampler with
pressures  ranging from 5 to  50  psi.  The  constant head tank enables the peristaltic
action  type pump to  operate with a  minimal variation in flow rate.  However, flow
rates and throughputs for the Model  LF-2  are from  10 to 25% lower than Model
LF-1 for parallel  sampling.
II.  PUMPING  SYSTEM

        Pumps, piping, and accessories are selected to. suit the specific conditions of
each station.  Shallow and deep well-type jet centrifugal pump systems are in use at
many stations to bring raw water from a representative sampling point to the sampling
apparatus.  Submersible pumps with helical screw rotors and synthetic rubber stators,
rotary pumps with flexible impellers, and other pumping mechanisms may  be installed
to meet individual needs.
        It is important that the pump does  not  contaminate the  sample through
grease-type packing or other sources.  The pump must have a greaseless-type rotary
shaft  seal or special packing material  to avoid  contamination.  Laboratory control
procedures  are  employed to assure that the pump does not contaminate  the sample.
New pumps are sometimes grease-coated and must be thoroughly cleaned before being
put into service.  Piping strainers,  check valves, and all other accessories that come in
contact with the raw water pumped to  the CAC  filter are also cleaned. (See Precau-
tions,  paragraph  below.)
III.  PRECAUTIONS

        The purpose of the CAC is to adsorb small  amounts of organic impurities
from the water in as great a quantity as possible. It is important to avoid contami-
nation of the carbon from other organic sources.  Hence the following precautions are
observed:

A. New strainers, pipe fittings, and other accessories  are usually coated with oil or
   grease.  The oil is removed by washing in kerosene or chloroform followed  by a
   detergent wash before fittings  are used for making connection to the  CAC.

B. Ordinary organic pipe jointing  compounds are not used.  Red lead  (lead oxide)
   mixed to a paste with water can be used  for this purpose.

                                      49

-------
C. Except as noted  below, plastic hose is avoided, and if rubber hose is used in any
   connections it is flushed thoroughly  before being connected to the CAC.  Copper
   tubing is  ideal for connections.  NOTE: Polyethylene pipe and PVC (polyvinyl
   chloride) pipe meeting National Sanitation Foundation (NSF) standards for drink-
   ing water use are acceptable.  Teflon  hose also is satisfactory for use.
IV.  USE  OF  CARBON  COLUMN  DATA SHEET

        Accurate flow  measurements are important. A sample data sheet is used to
record  flow  and other  pertinent information.
                                     50

-------
                       APPENDIX  TWO

      CHROMATOGRAMS, SAMPLE  CALIBRATION CURVES,
       INFRARED SPECTRA, AND STRUCTURAL  FORMULAE
  Heptochlor
        0   2   4   6   8   10  12
          A Retention Time in Minutes
    0    2   4   6   8   10  12
        B  Retention Time in Minutes
        Aldrin
              DDE
                      P,P' - DDT
                               O -o
                               O
   0    2   4   6   8   10  12   14
    C  Retention Time in Minutes
0  0.4 0.8  1.2 1.6  2.0  2.4  2.8  3.2
     D  PEAK AREA (Square Inches)
                              Figure 28

EC Gas Chromatograms of Standard Pesticides in (A) TLC Section II (he"ptachlor
epoxide—1 ng, dieldrin—2 ng, endrin—4 ng), (B) TLC Section III (lindane—1 ng,
y-chlordane—2 ng, ODD—3 ng), (C) TLC Section IV (heptachlor—0.5 ng, aldrin—
0.5  ng, DDE—1  ng, DDT—2  ng). (D) Sample Calibration  Curve for Dieldrin
(ECGC).  See page 28, footnote 30 for operating conditions.
                                 51

-------
     0   2   4   6   8   10  12  14
       A Retention Time in Minutes
                                    60

                                    50-
            ....  DDE
            Alarm  ,
     Heptachlor fl
                        p,p' - DDT
                    o/P-DDT
                                    10
0   24    6    8    10   12
   C   Retention Time in Minutes
                                       02   46   8   10  12
                                          B  Retention Time  in Minutes
                                               1        2        3
                                            D  PEAK AREA (Square  Inches)
                                   Figure 29

MCT Gas Chromatograms of Standard Pesticides in (A) TLC Section II (heptachlor
epoxide—25 ng, dieldrin—50 ng, endrin—100 ng), (B) TLC Section III (lindane—
20 ng, y-chlordane—40 ng, DDD—60 ng), (C) TLC Section IV (heptachlor—20 ng,
aldrin—20  ng,  DDE—40 ng, DDT—100 ng). (D) Sample Calibration Curve for
Endrin (MCTGC). See page 30, footnote 31 for operating  conditions.
                                      52

-------
 4000 3000      2000      1500
CM'      1000    900     800
                                                                    700
                            7     8     9     10     11
                             WAVELENGTH (MICRONS)
                      12    13
                                                                    14
SPECTRUM NO.
SAMPLE
DIBLDRIN STANDARD


QRIGIKlHirrRiTioNAL BIO-
CHEMICAL, CLEVELAND , CHIO
PURITY 85*
PHASF MINERAL OIL MULL
THICKNESS
i Fr;FKin
1
9
r>ATF W"
OPERATOR L.H.
REMARKS >"
%'
fn



      Figure 30.  IR Spectrum of Standard Dieldrin  in Mineral Oil Mull
 4000 3000
100
         1000   900     800
700
                            7     8     9     10    11
                             WAVELENGTH (MICRONS)
                                                         12    13    14    U
SPECTRUM NO.
SAMPLE
HffieiX STANDARD


OR'OIN NUTRITIONAL BIO-
Ct^MI^A1-- CLEVBLAND. OHIO
PI IPITY 9««
PHASF MINERAL OIL MULL
THICKNESS
IFftFWn
V
•)
nATF 7/7/M
OPERATOR L «-
RFMARKS >!
i!
m*



       Figure 31.  IR Spectrum of Standard Endrin in Mineral Oil Mull

                                     53

-------
                                     1000    900
800
                                                             700

                      7    8     9    10    11
                      WAVELENGTH (MICRONS)
                                                 12    13    14
SPECTRUM NO.
SAMPLE
LINDANB STANDARD


ORIGIN NUTRITIONAL BIO-
CHEMICAL. CLEVELAND. OHIO
PURITY REFERENCE STANDARD
PHASF MINERAL OIL MULL

IFrsFMD
1
9
HATF 7/7/64

RFMARKS >!
l!
m



Figure 32.  IR Spectrum of Standard Lindane in Mineral Oil Mull
                                     1000   900
                                             i,, ,
800
          700
                      7     8     ?     10    11     12    13    14    15
                       WAVELENGTH (MICRONS)
SPECTRUM NO.
SAMPLE
ODD STANDARD


ORIC5IN NUTRITIONAL BIO-
CHEMICAL, CLEVELAND, OHIO
PURITY REFERENCE STANDARD
PHASF MINERAL OIL MULL
THICKNESS
LEGEND

9
DATE '/7/64
OPFRATOR I-H.
RFMARKS




(/)
%
m

•

  Figure 33.   IR Spectrum of Standard ODD in Mineral Oil Mull

                               54

-------
              2000
                        1500
CM
                                                         800
                                                                  700
4000 3000
    3     4     5     6     7    8     9    10    11     12    13    14
                           WAVELENGTH (MICRONS)
SPECTRUM NO.
SAMPLE
HBPTACHLOR STANDARD


ORmiKlNUTRITIONAL BIO-
CHHMICAL. CLEVELAND. OHIO
PURITY 72*
PHASE MINERAL OIL HULL
THirKNESS
i Ff;Ft\(n
1

r>ATF 7/7/64
OPERATOR t.H.
REMARKS J
r
r



                                                                         •8

                                                                          o
    Figure 34.  IR Spectrum of Standard Heptachlor in Mineral Oil Mull
4000 3000
              2000
                           7     8     9     10    11
                            WAVELENGTH (MICRONS)
                                                       12     13    14    15
SPECTRUM NO.
SAMPLE
ALDRTN STANDARD


ORIGIN NUTRITIONAL BIO-
CHEMICAL, CLEVELAND, OHIO
PURITY «4"
PHASE MINERAL OIL MULL
THICKNESS
IFi^FKID
1
9
DATF 7A/64
OPERATOR >-. H.
REMARKS




i/>
X
MPIF



      Figure 35.  IR Spectrum of Standard Aldrin in Mineral Oil Mull

                                    55

-------
  4000 3000
 0.0
2000
  .10
 1500
i.i.
CM-'
1000    900
800
S.30
O
2-40

<50
 .60
 70

 1.0
                            ~7     8~    9    10
                              WAVELENGTH (MICRONS)
SPFCTRUM NO
SAAAPI F DDE
STANDARD


ORIOIN (EI
1
1 —



         Figure 37.  IR Spectrum of Standard DDT in Mineral Oil Mull


                                      56

-------
    ALDRIN
                                 Cl
Cl
    Cl     Cl


  HEPTACHLOR
                             ci
                                     ci
    ci

 LINDANE
                 Cl
                                                         Cl
HEPTACHLOR
  EPOXIDE
                                               Cl
       DDT
ODD
  DDE
Figure 38.  Structural Formulae of Nine Chlorinated Hydrocarbon Pesticides
                               57

-------
                               TABLE 1
Rf VALUES OF PESTICIDES DEVELOPED  WITH  CC14  ON  SILICA GEL-G
                         THIN LAYER PLATE
Pesticide
Methyl Parathion
Parathion
Dieldrin
Endrin
Heptachlor Epoxide
Lindane
DDD
y-Chlordane
Heptachlor"
DDT
DDE
Aldrin
Ri Value
0.05
0.07
0.17
0.20
0.29
0.37
0.54
0.55
0.67
0.68
0.72
0.73
Zone


II



III



IV

1 Technical grade heptachlor contains approximately 30% -x-chlordane.
                                  58

-------
                                                                TABLE 2
                                            GAS CHROMATOGRAPHIC  RETENTION  DATA
Temperature


Pesticide
Lindane
Methyl parathion
Heptachlor
Aldrin
Parathion
Heptachlor epoxide
y-Chlordane
Dieldrin
DDE (p, p')
DDD
DDT (o, p)
DDT (p, p1)
Endrin b
Electron Capture
180°C
Retention
Time In
Minutes
1.81
3.24
3.24
4.19
4.66
5.58
6.38
8.72
8.81
11.71
12.12
15.85
12.84-19.43
Relative a
Retention
Time
0.43
0.77
0.77
1.00
1.11
1.33
1.52
2.08
2.10
2.79
2.89
3.78
3.06-4.64
195°C
Retention
Time In
Minutes
0.94
1.58
2.03
—
2.61
3.00
3.87
3.94
4.96
5.20
6.62
5.20-7.40
Relative "
Retention
Time
0.46
0.78
1.00
—
1.29
1.48
1.91
1.94
2.44
2.56
3.26
2.56-3.64
Microcoulometric
190°C
Retention
Time In
Minutes
1.90
3.07
3.80
—
4.72
5.40
6.95
6.79
8.87
9.09
11.39
9.63-13.65
Relative "
Retention
Time
0.50
0.81
1.00
—
1.24
1.42
1.83
1.79
2.33
2.39
2.99
2.53-3.59
(O
          " Ratio of absolute retention time for compound to that of aldrin.
          "The multiple peaks for endrin have been associated with the thermal isomerization of endrin on gas chromatographic columns (63).

-------
                                 TABLE 3
    SOME COLUMN PACKINGS USED FOR GAS  CHROMATOGRAPHIC
       ANALYSIS OF CHLORINATED HYDROCARBON PESTICIDES
      Solid Support
Chromosorb  W (60-80 mesh)
Chromosorb  W (60-80 mesh)
Chromosorb  W (60-80 mesh)

Chromosorb W (60-80 mesh) acid washed
Chromosorb W (60-80 mesh)
  silanized (HMDS)
Chromosorb W (100-120 mesh)
  silanized (HMDS)
Chromosorb W (45-60 mesh)
  silanized (HMDS)
Chromosorb P (60-80 mesh) acid washed
Chromosorb P (30-60 mesh) acid washed

Chromosorb P (30-60 mesh) acid washed
Chromosorb P (30-60 mesh) acid washed
Chromosorb P (60-80 mesh) acid washed
Fluoropak-80 preceded by
Anakrom ABS (90-100 mesh)
Gas Chrom Q  (100-120 mesh)
Liquid Phase               Reference
5% Dow 11 Silicone            (14)
10% SE-30 Silicone Gum       (45)
5% FS-1265 (QF-1)  Fluoro     (21)
  Silicone
4% SE-30 Silicone Gum +     (64)
  6% FS-1265 (QF-1)
  Fluoro Silicone
2.5% Dow 11 Silicone         (65)

5%Apiezon-L                 (20)

5% SE-30 Silicone Gum       (20)

5% DC-200 Silicone            (46)
15 to 30% High Vacuum       (52)
  Silicone Grease              (66)
20% SE-30 Silicone Gum       (67)
2.5% Epon Resin 1001         (68)
5% FS-1265 (QF-1)  Fluoro     (69)
  Silicone -f 3% DC-200
  Silicone
12% Dow 11 Silicone  +       (47)
  6%  Epon 1001
10% DC-200 Silicone          (70)
    DC-200 Silicone           (71)
                                     60

-------
                       APPENDIX  THREE
             EQUIPMENT, SOLVENTS  AND REAGENTS


I.  EQUIPMENT,  SOLVENTS  AND  REAGENTS  USED  TO COLLECT
   AND PROCESS CCE  SAMPLES  (PER  SAMPLE)

A.  EQUIPMENT
Collection equipment  (see Appendix One.)
 1  Drying oven.
 1  1-gallon paint can with crimp lid.
 1  Extraction assembly, includes:
       a. 2  3-liter round bottom flasks
       b. 1  large Soxhlet extractor
       c. 1  Friedericks condenser
       d. 1  3-liter heating mantel
             (Glas-Col Series M)
       e. 1  Variable transformer
             (Powerstat, Type 116)
 1  Manifold for filtering and warming air for drying carbon in Soxhlet.
 1  10-ml beaker.
 1  50-ml beaker.
 1  100-ml beaker.
 2  300-ml Erlenmeyer flasks.
12  125-ml Erlenmeyer flasks.
 1  125-ml vacuum flask.
 1  60°-sintered-glass funnel.
11  5-dram glass vials.
 1  chromatographic column (19 mm I.D. with  medium porosity fritted disc).
 1  100-ml graduated cylinder.
 1  each 10-/J, 50-fil and  100-/nl  syringes for gas chromatographic injections.

B. SOLVENTS (all redistilled  A.R. grade)
3 liters chloroform
3 liters ethanol (95%)
100 ml of methanol
100 ml of isooctane
100 ml of benzene
300 ml of ethyl ether

                                  61

-------
C.  REAGENTS
50 ml HC1 (cone.)
50 ml HC1 (5%)
50 ml NaHCOa (5%)
50 ml NaOH (5%)
NaOH (25%) or pellets
20 grains activated silica gel (Davison Code 950-08-08-226, 60-200 mesh)

D.  PESTICIDE  STANDARDS


II.  EQUIPMENT, SOLVENTS AND  REAGENTS  USED  TO  COLLECT
    AND  PROCESS WATER GRAB  SAMPLES  (PER  SAMPLE)

A.  EQUIPMENT
1  1-quart glass bottle with Teflon liner in cap.
1  Constant temperature water bath  (40°C).
   Compressed air source (clean-dry).

1.  Semi-Automatic Extraction
   1  Semi-automatic extractor includes:
        1  Teflon impeller containing bar magnet (Teflon is used because it does not
        contaminate the solvents and is highly resistant to corrosive cleaning agents).
        1  Bottle lid with the  center removed and a double Teflon liner inserted.
   1  Magnetic stirrer.
   1  Teflon reclamation plug with glass recovery tube.
   1  Pressure device  to secure reclamation plug.
   1  Source of compressed air.
   1  50-ml graduated cylinder.
   1  100-ml beaker.
   1  1000-ml  graduated cylinder.
 • 1  15-ml graduated centrifuge tube.

2.  Separately Funnel Extraction
   1  2-liter separatory funnel  with Teflon stopcock.
   1  1500-ml beaker.
   2  300-ml Erlenmeyer flasks.
   1  Chromatographic column (19 mm I.D.).
   1  each, 10-/xl,  50-/*1, and 100-^1 syringes for gas chromatographic injections.
   1  Kuderna-Danish Evaporator with ampoule graduated in 0.01 ml divisions.

B.   SOLVENTS (CHROMATOGRAPHIC GRADE)
1.  Semi-automatic Extraction
   50 ml hexane-benzene (1:1)
2.  Separatory Funnel Extraction
   300 ml of hexane.

C.  REAGENTS
Anhydrous sodium sulfate.

                                    62

-------
III.   EQUIPMENT,  SOLVENTS AND  REAGENTS  USED TO  PROCESS
     BOTTOM  SAMPLES
Each extraction apparatus includes:
1  300 ml round bottom flask T 24/40.
1  Soxhlet extractor, 40 mm I.D. x 205 mm length, bottom joint f  24/40, top joint
   T 45/50.
1  Allihn condenser joint T 45/50
   Pre-extracted glass wool.
1  Heating mantle, 300 ml.
   200 ml of hexane-acetone (9:1).
1  Large porcelain mortar and pestle.

IV.  EQUIPMENT,  SOLVENTS  AND  REAGENTS  FOR  THIN  LAYER
     CHROMATOGRAPHY
Plates, 200 mm x 200 mm (8" x 8") glass
Chamber, glass  developing, with lid,  Sl/2r' x 4" x S1/^"
Spreader, variable thickness
Thickness gauge
Plate holder, plastic
Plate carrier
Spotting template
Chromatography sprayer
Desiccator to accommodate 200-mm plates
UV light box
Micropipets, 1/*1—10 /J, 100 /*!
Eye droppers
Spatulas
Graduated centrifuge  test  tubes, capacity 15 ml
Glass wool, pre-extracted with  chloroform
Pesticide standards
  All pesticides 0.1% w/v  (1  mg/ml) in hexane
Chromogenic agents:
  Rhodamine B base, spirit soluble 0.10 mg/ml in ethanol
  Silver nitrate solution
    1.7 g AgNOs in 5 ml distilled water is  added to 10 ml 2-phenoxyethanol (MC &
    B, technical), and diluted to  200 ml with  acetone.
  5% bromine in carbon tetrachloride
    Fluorescein solution
    MC & B fluorescein, water soluble, U.S.P.,  25 mg. in 100 ml dimethylformamide;
    10 ml of this concentrated solution are diluted to  50 ml with ethanol  95%  for
    spraying.
Adsorbents:
  Silica Gel G
  Aluminum Oxide G
Solvents:
  All solvents are redistilled before use
  Hexane
  Carbon Tetrachloride
  Ethyl ether-petroleum ether  (1:1)
  Acetone
Dye Mixture:
  Sudan Yellow, Sudan IV, Azobenzene, 0.1% in benzene

                                     63

-------
                          APPENDIX  FOUR


  GENERAL  COMPOSITION  OF CARBON CHLOROFORM AND

                    CARBON  ALCOHOL  EXTRACTS


I.  CHLOROFORM  EXTRACTS

        The organic residue recovered  from the carbon adsorption column by chloro-
form is very complex.  It is  desirable to separate the crude extract into certain broad
chemical  classes, and this can be done on the basis of solubility differences. The var-
ious classes or groups and their general significance  are discussed briefly below.
          ///
A.  ETHER.SOLUBLES
          /i
        This group is usually a brown, humus-like powder, apparently composed to
a large extent of carboxylic acids, ketones,  and  alcohols of complicated  structure.
Origin of the group, which is an indicator of "old" pollution, is believed to be partially
oxidized sewage and industrial wastes.  For example, the Ohio River at Cincinnati has
been exposed to much industrial and sewage  pollution, and  hence large amounts of
ether-insoluble materials are found.  Streams with little or no pollution history have
little or no ether insolubles. Chloroform extracts contain from 0 to 30  percent of
ether-insoluble material.



        These substances are largely acidic  and  undistillable at moderate tempera-
tures, but their solubility in ether indicates that the molecules are smaller and prob-
ably simpler than the  ethewsolubles.  On the  other  hand, their water  solubility
practically requires the presence  of  several  functional groups, such as hydroxy-acid,
keto-acid, and keto-alcohol. Such  compounds probably originate from partial oxida-
tion of hydrocarbons or they may  be natural substances.  They  have very little odor.
These materials usually make up 10  to 20 percent of the  total extract.

C.  WEAK ACIDS
        This group is  characterized by being removed from ether solution with sodium
hydroxide but not with sodium bicarbonate.  Phenols are  the best known weak acids,
and if present in the water, appear in this group.  Other weakly acidic compounds
include certain enols, imides, sulfonamides, and some sulfur compounds. This group of
materials also occurs in nature. The weak acids are odorous, and commonly constitute
5 to 20 percent of the chloroform extract.

                                     64

-------
D.  STRONG ACIDS
        These  acids  are usually carboxylic acids such as acetic, benzoic, salicylic or
butyric. Although classified as strong  in reference to carbonic acid, they are actually
weak when compared with a mineral acid, such as sulfuric.  Many of the compounds
are used industrially, but may also be produced by natural processes, such as fermen-
tation. Some of the materials are highly odorous.  This fraction makes up from 5 to
20 percent of the total. The significance of the strong acids can be interpreted  only
in the light of stream pollution conditions.


E.  BASES
        These compounds are organic amines.  Such materials as aniline and pyridine
are amines of commerce.  Lower  amines may occur  as  a  result  of decomposition.
Although odorous, the lower concentrations found are not  likely to cause objectionable
conditions. However, in the case of  specific  amine-containing  wastes  the compounds
can be of considerable significance.  Generally, only 1 or 2 percent of the total extract
is made up of the bases.


F.  NEUTRALS
        This group  frequently constitutes the major portion of the chloroform extract.
Neither basic nor acidic, the materials are less reactive and tend to persist in streams
longer than many other types. Hydrocarbons, aldehydes, ketones, esters, and ethers are
examples  of neutral  materials.  The group  lends itself  to  further fractionation  by
chromatographic  separation into  aliphatic, aromatic, and oxygenated  subgroups:


1.  Aliphatics:
        This portion represents petroleum-type hydrocarbons in a considerable  state
of purity, and is usually made up of mineral oil type of  material.  The percentage of
aliphatics  present yields important information about the possible source of pollution,
since  petroleum is the most likely source.


2.  Aromatics:
        These are principally the coal tar hydrocarbons such as benzene, toluene, and
a host of others, and their presence in any significant amount is a  reliable indication
of industrial pollution. Further, the materials can frequently be identified by infrared
spectrophotometry. Some aromatic  compounds which have been found in our rivers—
and in our drinking water—include  DDT, aldrin, dieldrin, endrin, phenyl ether, ortho-
nitrochlorobenzene, pyridine, phenol, and others.  Some of these materials are highly
odorous; others may also be toxic.  Their appearance in  any  quantity  as pollutants
should receive careful evaluation.
3.  Oxygenated Compounds (Oxys):
        These are the neutral compounds containing oxygen  in  aldehyde, ketone, or
esters groups.  They may have originated  by direct discharge or may represent oxida-
tion products  from both natural and industrial materials.  They help to indicate the
"age"  of  the  pollution, since pollution  exposed  to oxidation forces  for a long time
would be expected to contain large amounts of oxys.  The oxy materials are odorous.

                                        65

-------
G.   LOSSES
         Manipulative losses inherent in this type of separation may amount to 10 to
15 percent.  Losses greater than this may indicate that volatile components were lost.
from the sample.  Such volatile.? may have significance as pollutants.


II.   ALCOHOL  EXTRACTS

         The alcohol extractables generally consist of materials more polar than the
chloroform extractables. They often contain  synthetic detergents, carboxylic acids and
humic  materials which may  originate naturally or from oxidized products of domestic
and  industrial wastes.  These classes of substances  are not quantitatively recovered
by the alcohol extraction.  For example, this extraction recovers only 20 to  30 percent
of the  synthetic detergents present.  On waters of mixed industrial and domestic pollu-
tion, the chloroform and alcohol extractables may  be about equal.  On some streams
where  the industrial pollution is rather  low and much  natural pollution or sewage is
present, the alcohol extractables  may exceed the chloroform extractables by a factor
of 4  to 6.
        The alcohol  extract is usually  only partially soluble in water and  most ordi-
nary solvents.  Very  little further chemical separation of this material is currently
practical.  However, tests have revealed that synthetic detergents may make up 1 to
12 percent of the alcohol extract.
                                       66

-------
                           APPENDIX  FIVE
                                GLOSSARY
mg       milligram  (10~3 gram)
/*g        microgram (10~6 gram)
ng        nanogram  (10~9 gram)
pg        picogram  (10~12 gram)
ml        milliliter (1Q-3 liter)
/xl        microliter  (10~6 liter)
CAM     carbon adsorption method
CAC      carbon adsorption column
CCE      carbon chloroform extract
CAE      carbon alcohol extract
El        ether insolubles
WS       water solubles
B        bases
SA        strong acids
WA       weak acids
N        neutrals
AL       aliphatics
AR       aromatics
OXY     oxygenated substances
IR       infrared
TLC      thin layer  chromatography
GC       gas chromatography
ECGC    electron capture gas
            chromatography
MCTGC  microcoulometric titration gas
            chromatography

          distance travelled by a given
            substance
Rf        	
          distance travelled by solvent
            front
                                REFERENCES
1. Braus,  H., Middleton, F.  M., and
Walton,  G.,  Anal.  Chem.,  23,   1160
(1951).

2. Middleton, F.  M., Grant,  W., and
Rosen, A. A., Ind. Eng.  Chem., 48, 268
(1956).

3. Middleton, F. M., and Rosen, A. A.,
Public Health Reports, 71, 1125 (1956).

4. Ludzack, F. J., Middleton, F. M., and
Ettinger, M. B., Sewage  & Ind. Wastes,
30, 662 (1958).

5. Palange, R.  C.,  and  Megregian, S.,
J.A.W.W.A.,50, 1214 (1958).
6.  Palange,  R.  C.,  and Megregian,  S.,
Proc.  A.S.C.E., 84-.SA2, Paper No. 1606
(1958).

7.  Middleton, F.  M., and  Lichtenberg,
J. J., Ind. Eng. Chem., 52, 99A (1960).

8.  Hoadley, A. W.,  "A Preliminary Re-
view  of Carbon  Adsorption Data Col-
lected  as Part  of the National Water
Quality Network  Sampling  Program."
National Water Quality Netwerk Appli-
cations  and Development  Report #5,
Public Health Service, Division of Water
Supply  and  Pollution  Control,  Basic
Data  Branch, Water Quality  Section,
Cincinnati,  Ohio, May 1962.
                                      67

-------
9.  Booth, R. L., English, J. N., and Mc-
Dermott, G.  N., J.A.W.W.A., 57, 215
(1965).

10. Booth,  R.  L., "Optimum Sampling
Rate and Sample Volume for Quantita-
tive Measurement of  Organics by the
Present  Carbon  Adsorption  Method,"
Department of Health, Education, and
Welfare, Public Health Service, Division
of Water Supply  and Pollution Control,
Robert A. Taft  Sanitary  Engineering
Center,  Cincinnati,  Ohio,  August  26,
1963.


11. Lee,  G. F., et al., Int. J. Air Water
Pott., 9, 69 (1965).
12.  Greenberg, A. E., Maehler, C. Z., and
Cornelius,   J.,  J.A.W.W.A.,  57,  791
(1965).
13.  Rosen, A. A., and Middleton, F. M.,
Anal. Chem., 31,1729 (1959).


14.  Breidenbach, A.  W.,  and  Lichten-
berg, J. J., Science, 141, 899 (1963).
15. Nicholson,  H.  P.,  et  al., Limnol.
Oceanog., 9, 310 (1964).
16. Grzenda,  A. R.,  et  al.,  J. Econ.
Entomol., 57, 615 (1964).


17. Goodenkauf,  A.,  and  Erdei,  J.,
J.A.W.W.A., 56, 600  (1964).
20. Kahn, L., and Wayman, C. H., Anal.
Chem., 36, 1340 (1964).

21. Lamar, W. L., Goerlitz, D.  F., Law,
L. M., "Determination and Measurement
of  Chlorinated  Organic  Pesticides in
Water by Electron Capture Gas Chro-
matography,"  Open-File Report, U. S.
Dept. of the Interior, Geological Survey,
Water  Resources  Division,  November
1964.

22. Berck,  B., Anal.  Chem.,  25, 1253
(1953).

23. Lichtenstein,  E. P., J. Econ.  En-
tomol., 50, 545 (1957).

24. Lichtenstein, E. P.,  and Schultz, K.
R., ibid., 54, 517 (1961).

25. Gannon, N., and Bigger, J.  H., ibid.,
51, 1  (1958).
26. Wilkinson, A. T. S.,  and Finlayson,
D. G., Science, 143, 681 (1964).


27. Middleton, F. M., Rosen, A. A., and
Burttschell, R. H., "Manual for  the Re-
covery  and  Identification  of  Organic
Chemicals  in  Water," Rob'ret A. Taft
Sanitary Engineering Center, Cincinnati,
Ohio, May, 1959.

28. Middleton, F. M., Greenberg, A. E.,
and  Lee,  G.  F., J.A.W.W.A.,  54, 223
(1962).
18.  Breidenbach, A. W., et al., "Chlori-
nated Hydrocarbon Pesticides in Major
River Basins, 1957-1965,"  Presented  at
the Joint Meeting of  the Commissioned
Officers   Association   and  the  Clinical
Society  at  Baltimore, Maryland, 1966.
29. Booth, R.  L.,  "Reproducibility  of
Carbon Adsorption  Method  (CAM),"
Memo to Sub-Committee on Applicabil-
ity of Carbon Adsorption Technique  to
the Mission of the National Water Qual-
ity Network, January 31, 1963.
19. Weaver, L., et al., Public Health Re-     30. Castelli,  J.  A.,  and Booth,  R. L.,
ports, 80, 481  (1965).                      J.A.W.W.A.,  56, 1243 (1964).
                                       68

-------
31.  Reid,  B.  H.,  et al., "Preliminary
Field Evaluation of Low Flow Rate Car-
bon Adsorption Equipment and Methods
for   Organic   Sampling   of   Surface
Waters," PHS Water Pollution Surveil-
lance System Application and Develop-
ment Report  #14,  Division  of  Water
Supply  and  Pollution  Control, Basic
Data Branch,  Water  Quality Section,
Cincinnati, Ohio, March 1965.
32. "A Study of Methods Used in Meas-
urement and Analysis of Sediment Loads
in Streams, Report  M—Operation and
Maintenance of U. S. BM-54 Bed-Mate-
rial  Sampler,"  St. Anthony  Falls  Hy-
draulic  Laboratory,  Minneapolis,  Min-
nesota, November 1958.
33. Mashni,  C.,  Private  Communica-
tion,  Federal  Water  Pollution  Control
Administration, Cincinnati, Ohio, August
1963.
34. Shriner, R.  L., Fuson,  R, C., and
Curtin,  D. Y.,  "The Systematic Identi-
fication  of  Organic  Compounds," 4th
Edition, John Wiley & Sons, New York,
N.Y., 1956.
35. Cheronis, N. D., and Entrikin, J. B.,
"Semimicro  Qualitative  Organic Anal-
ysis,"  Thomas  Y. Crowell  Co., New
York, N.Y.,  1947.
36.  Cheronis, N. D., "Micro  and Semi-
micro Methods," Vol. VI of  the  Series,
"Techniques  of  Organic  Chemistry."
Arnold Weissburger, Editor, Interscience,
New York, N.Y., 1954.
       39.  Mills,  P.  A., J.A.O.A.C.,  42, 734
       (1959).

       40.  Johnson,  L.,  J.A.O.A.C.,  45, 363
       (1962).


       41.  Mitchell, L. C., J.A.O.A.C., 41, 781
       (1958).

       42.  Walker,  K.  C.,  and  Beroza, M.,
       J.A.O.A.C., 46,250 (1963).


       43.  Reid,  B. H.,  Patent Applied For,
       "Semi-Automatic  System  for  Solvent
       Extraction of Organic Substances from
       AVater,"  DWSPC  Case  No.  E-65-11,
       March  12,  1965.   Present   Address:
       FWPCA,  Pacific Northwest Water Lab-
       oratory, Corvallis, Oregon.


       44.  Kawahara,  F. K.,  et  al., "Semi-
       Automatic Extraction of Organic Mate-
       rials from Water," PHS Water  Pollution
       Surveillance  System  Applications and
       Development Report #16, Division  of
       Water Supply  and Pollution  Control,
       Basic Data Branch, Water Quality Sec-
       tion, Cincinnati, Ohio, November 1965.


       45.  Julian, E. C., Private Communica-
       tion,  Federal  Water  Pollution Control
       Administration,  Cincinnati, Ohio, May
       1963.


       46.  Teasley,  J.  I.,  and Cox, W.   S.,
       J.A.W.W.A., 55, 1093  (1963).


       47.  Schafer,  M.,  Busch,  K.  A.,  and
       Campbell, J. E., J. Dairy Science, XLVI,
       1025  (1963).
37.  Schneider, F.  L.,  "Qualitative Or-
ganic  Microanalysis,"  John  Wiley  &
Sons, New York, N.Y., 1946.
       48. Rosen, A. A., Private Communica-
       tion,  Federal  Water Pollution  Control
       Administration, Cincinnati, Ohio, 1964.
38.  Smith,   D.  and  Eichelberger,
J.W.P.C.F., 37, 77 (1965).
J.,     49.  Shell Development Co., Agricultural
       Research Division,  Modesto, California,
                                       69

-------
"Extraction of Halogenated Hydrocar-
bon Pesticide Residues from Crops, Soils,
and Animal Products," Analytical Meth-
ods, MNS—1/63,1963.

50. Burchfield, H. P., and Schuldt, P. H.,
Contrib. Boyce Thompson Inst., 19, 77
(1957).

51. Lovelock,  J. E., and Lipsky, S. R.,
J. Am. Chem. Soc., 82, 431  (1960).

52. Coulson, D. M., et  al., J. Agr. and
Food  Chem., 8, 399 (1960).

53. Karmen, A., Anal. Chem.,  36,  1416
(1964).  ,

54. Giuffrida,   L., J.A.O.A.C.,  47, 293
(1964).

55. Micro-Tek Instruments, Inc., Baton
Rouge,  Louisiana, "Tek-Talk,"  Volume
1, Number 2, Summer 1965. '

56. Brody, S.  S. and Chaney, J. E.,  J.
Gas Chromatog., 4, 42 (1966).

57. Bache,  C.  A., and Lisk, D. J., Anal.
Chem., 37,  1477 (1965).

58. Burchfield, H.  P.,  et  al.,  J.  Gas
Chromatog., 3, 28 (1965).

59. Wren,  J. J., J. Chromatog.,  4, 173
(1960).

60. Moddes,  R.,  J.A.O.A.C.,  44,  169
(1961).

61. Kovacs, M. F., J.A.O.A.C.,  46, 884
(1963).

62. U. S. Department of Health, Edu-
cation,  and Welfare, Food  and  Drug
Administration,  Pesticide   Analytical
Manual, Volume I, Chlorinated 2.34 (b),
"Thin Layer Chromatography for Chlo-
rinated   Pesticide  Residue  Analysis,"
July 1964.

63.  Phillips, D. D., Pollard, G. E., and
Soloway, S. B., J. Agr. and Food Chem.,
10,  217  (1962).

64.  McCully, K. A. and Mckinley, W. P.,
J.A.O.A.C., 47, 652  (1964).

65.  Simmons, W.  H., Private Communi-
cation,  Woodson-Tenent   Laboratories,
Memphis, Tennessee (1965).

66.  Burke, J.  and Johnson, L.,  J.A.O.-
A.C., 45, 348 (1962).

67.  Cassil, C.  C.,  "Pesticide  Residue
Analysis  by  Microcoulometric   Gas
Chromatography,"  Residue   Reviews,
Volume  I, edited  by F.  A. Gunther,
Springer-Verlag, Berlin, Gottingen, Hei-
delberg, Germany, 1962.

68.  Goodwin,  E.  S., Goulden,  R.,  and
Reynolds, J.  G.,  "Gas Chromatography
with Electron Capture  lonization Detec-
tion for  Rapid Identification of  Pesti-
cide Residues  in  Crops,"  Eighteenth
International Congress of  Pure and Ap-
plied Chemistry, Montreal, August 1961.

69.  White, R. E., "Insecticide Analysis
Procedures  Used  by  Klamath  Basin
Study," Presented at the Pacific North-
west Pollution Control  Association, Van-
couver,   British   Columbia,  November
1965.

70.  Shuman,  H.,  and Collie,  J.   R.,
J.A.O.A.C., 46, 992  (1963).

71.  Applied  Science Laboratories, Inc.,
State  College,   Pennsylvania,   "Gas-
Chrom Newsletter," December 1965.
          Mention of products and manufacturer is for identification only
                      and does not imply endorsement by the
                 Federal Water  Pollution Control Administration
                      or the U.S. Department of the Interior
                                         •fr U.S. GOVERNMENT PRINTING OFFICE: I96S  O22O-916
                                       70

-------