Development and Evaluation of Analytical
Test Procedures for Priority Pollutants
Gulf South Research Inst.
New Orleans, LA
Prepared for
Environmental Monitcring and Support
Lab.-Cincinnati, OH
Feb 83
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P1363-166181
EPA-600/^-83-002
February 1983
DEVELOPMENT AND EVALUATION OF ANALYTICAL
~TEST PROCEDURES FOR PRIORITY POLLUTANTS
!
\>y
C. S. Monteith
Gulf South Research Institute
New Orleans, Louisiana 70186
Contract No. 68-03-2779
Project Officer
D. F. Bender
Environmental Monitoring and Support Laboratory
U.S. Environmental Protection Agency
Cincinnati, Ohio 45268
OFFICE OF RESEARCH AND DEVELOPMENT
U.S. ENVIRONMENTAL PROTECTION AGENCY
CINCINNATI, OHIO 45268
KPRODUCED BY
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TECHNICAL REPORT DATA
(Please read /atuvctions on the reverse before eompletingj
1. REPORT NO.
EPA-600/'»-83-002
2.
3-Rwrcc
ESSIOf*NO.
16618 1
4. TITLE AND SUBTITLE
Development and Evaluation of Analytical Test Pro-
cedures for Priority Pollutants
5. REPORT DATE
February 1983
6. PERFORMING ORGANIZATION CODE
7. AUTHOR(S)
C. S. Monteith
B. PERFORMING ORGANIZATION REPORT NO.
1 HD 621
9. PERFORMING ORGANIZATION NAME AND ADDRESS
Gulf South Research Institute
New Orleans, Louisana 70186
10. PROGRAM ELEMENT NO.
11. CONTRACT/GRANT NO.
Contract Number 68-03-2779
12. SPONSORING AGENCY NAME AND ADDRESS
Environmental Monitoring and Support Laboratory
13. TYPE OF REPORT AND PERIOD COVERED
Office of Research and Development
U.S. Environmental Protection Agency
Cincinnati, Ohio 45268
14. SPONSORING AGENCY CODE
EP/1/S00/06
15. SUPPLEMENTARY NOTES
16. ABSTRACT
Analytical methods were developed for the determination of cyanide and of
total phenolic compounds in solid/semi-solid samples of environmental
importance. A number of methods were reviewed, selected methods were
empirically evaluated,and the most promising methods were optimized and
validated by application to ten solid/semi-solid matrices. Each method
consisted of two steps: isolation of the analyte and quantification of the
analyte. Isolation studies were performed using radiolabeled cyanide and
radiolabeled phenolic compounds to spike representative matrices.
The method which was selected for the determination of cyanide in
solid/semi-solid matrices involves distillation of cyanide from, an acidified
slurry of the sample, followed by quantification using the pyridine-barbituric
acid procedure. The method which was selected for the determination of total
phenolic compounds involves extraction of an acidified slurry of the sample
with methylene chloride in a blender. The methylene chloride is then
extracted with a sodium hydroxide solution and quantified using the
4-aminoantipyrene (4-AAP) procedure.
17.
KEY WORDS AND DOCUMENT ANALYSIS
a. DESCRIPTORS
b.lOENTIFIERS/OPEN ENOED TERMS
c. COSATI Ficld/Gioup
18. DISTRIBUTION STATEMENT
release to public
19. SECURITY CLASS (This Report)
unclassified
21. NO. OF PAGES
103
20. SECURITY CLASS (Thispage)
unclassified
22. PRICE
EPA Form 2220-1 (9*73) ]
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NOTICE
This document has been reviewed in accordance with
U.S. Environmental Protection Agency policy and
approved for publication. Mention of trade names
or commercial products does not constitute endorse-
ment or recommendation for use.
ii
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FOREWORD
Environmental measurements are required to determine the quality ot
ambient waters and the character of waste effluents. The Environmental
Monitoring and Support Laboratory - Cincinnati conducts research to:
i
• Develop and evaluate methods to measure the presence and concentra-
tion of physical, chemical, and radiological pollutants in water,
wastewater, bottom sediments, and solid waste.
Investigate methods for the concentration, recovery, and identifi-
cation o£ viruses, bacteria and other microbiological organisms in
water; and, to determine the responses of aquatic organisms to
water quality.
* Develop and operate an Agency-wide quality assurance program to
assure standardization and quality control of systems for monitor-
ing water and wastewater.
Develop and operate a computerized system for instrument auto-
mation leading to improved data collection, analysis, and quality
control.
This report describes the evaluation of methods for the analysis of cyauide
compounds and phenolic compounds in ten solid and semisolid matrices which
are of environmental importance. The methods were optimized primarily on
bottom sediments. These data, and the conclusions which can be drawn from
them, will be an important part of future decisions and future recommeda-
tions concerning methods of analysis for solids and semisolid samples which
are related to the pollution of our water sources.
Robert L. Booth, Acting Director
Environmental Monitoring and Support
Laboratory - Cincinnati
iii
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ABSTRACT
The goals of this research program were to develop analytical methods
; for cyanide and total phenolic compounds in solid and semisolid environ-
i mental samples. The program was divided into four phases: review of the
• literature and selection of methods, empirical evaluation of methods, opti-
: mization of promising methods, and validation of the optimized methods.
Procedures for isolation and quantification were evaluated independently.
Isolation procedures were evaluated using solid and semisolid environmental
samples spiked with radiolabeled cyanide or phenolic compounds. Quantifi-
. cation procedures were tested initially on aqueous solutions of cyanide or
. phenol. Promising isolation and quantification procedures for cyanide or
¦' phenolic compounds were united to form a single method for analysis of solid
and semisolid samples. These methods were optimized and validated by analy-
sis of samples of ten solid and semisolid environmental matrices. Preserva-
tion procedures for these samples were also investigated.
The results of this research effort are methods for analysis of solid
; and semisolid environmental samples for cyanide and total phenolic com-
i pounds. The method for determination of cyanide is distillation of cyanide
, from an acidified slurry of the sample followed by quantification using the
: pyridine-barbituric acid procedure. Phenolic compounds are extracted from
' acidified slurries of solid and semisolid samples by blending with methylene
chloride. The phenolic compounds are extracted from the methylene chloride
into sodium hydroxide solution and quantified using 4-aminoantipyrine solut-
| ion.
This report was submitted in fulfillment of Contract No. 68-03-2779 by
Gulf South Research Institute under the sponsorship of the U. S. Environmen-
tal Protection Agency, and covers a period from February 12, 1979, to
February 11, 1981, and work was completed as of February 11, 1981.
Iv
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CONTENTS
Page
Foreword ; iii
Abstract ! iv
Figures vi
Tables .' vi
1. Introduction 1
2. Recommendations and Conclusions 3
3. Method for Analysis of Solid and Semisolid Environmental
Samples for Cyanide 4
Introduction - 4 .
Phase I: Selection of Methodology from the-Literature.. 5
Phase II: Evaluation of the Methods 15
Phase III: Optimization of Method 24
Phase IV: Validation of Method 28
4. Method for Analysis of Solid and Semisolid Environmental
Samples for Total Phenolic Compounds 33
Introduction 33
Phase I: Selection of Methodology from the Literature.. 34
iPhase II: Evaluation of the Methods 38
Phase III: Optimization of the Method 47
Phase IV: Validation of the Method 50
5. Preservation of Cyanide and Phenolic Compounds in Solid
and Semisolid Environmental Samples 56
Introduction 56
Methods of Preservation 57
Evaluation of Preservation Methods 57
Validation of Preservation Methods 59
References 62
Appendices
A. Procedures for Quantification of Cyanide 74
B. Method for Analysis of Solid and Semisolid Environmental
Samples for Cyanide, Total 87
C. Procedures for Quantification of Total Phenolic Compounds 92
D. Method for Analysis of Solid and Semisolid Environmental
Samples for Total Phenolic Compounds 99
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FIGURES
lumber rage
1 Apparatus for distillation of cyanide 8
2 Cyanide distillation apparatus equipped with a lead
acetate-filled absorber tube 26
.TABLES
Jumber , Page
1 Comparison of Recovery of Cyanide from Sediment Held
0.5 hr and 16 hr before Distilling 18
2 Percentages and Standard Deviations of Cyanide Recovered
from Three Matrices by Distillation Procedures 19
3 Recovery of Cyanide from Fish Tissue, Sediment and Sludge by
Distillation with Several Refluxing Reagents 21
4 Results of Evaluation of Procedures for Determination
of Cyanide 23
5 Percent Recovery of Cyanide by Solvent Extraction as
Cyanogen Bromide 24
I
6 Comparison of Percentages of Cyanide Recovered from Fish Tissue
Spiked with 100 Micrograms of KCN.. 27
\
7 Micrograms of Cyanide Recovered from Spiked and Unspiked
Sample of Fish Tissue 28
' *
8 Concentrations of Cyanide Added to and Recovered from
Solid and Semisolid Environmental Samples 30
9 Recovery of Cyanide from Aqueous Quality Assurance Samples 32
10 Average Percent Phenol Extracted from Sludge by the Stir-Bar
Extraction Procedure without Volume Replacement 41
; I
.11 Average Percent Recovery of Phenolic Compounds 42
vi'
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Number Pafte
14
12 Recovery of C-Labeled Phenolic Compounds from Sludge,
Sediment, and Fish by Soxhlet Extraction 43
J
—13 Average Percent Recovery of Phenol, P-Nitrophenol, and
-2,4,5-Trichlorophenol by Distillation -44
14 Range, Precision, and Accuracy of Procedures for the
Determination of Total Phenolic Compounds 46
15 Average Percentages and Standard Deviations of Phenol,
Nitrophenol, and Trichlorophenol 49
16 Micrograms of Phenol Recovered from Spiked and Unspiked
Samples of Sludge 51
17 Concentrations of Phenol Added to and Recovered from Solid
and Semisolid Environmental Samples 53
18 Recovery of Phenol from Aqueous Quality Assurance Samples 55
14
19 Percent Recovery of C-Labeled Cyanide from Water Samples 58
14
20 Percent Recovery of C-Labeled Phenol from Samples of
Sludge and Water 60
21 Concentrations of Cyanide Cpg/g) in Sludge Samples 60
22 Concentrations of Phenol (jJg/g) in Soil Samples 61
i
vil
iVJf.'t
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SECTION 1
INTRODUCTION
Cyanide and phenolic compounds are common pollutants of water bodies
and land disposal sites. Cyanide complexes are found in the wastewaters of
many industrial processes, including electroplating, precious metal refin-
ing, and steel hardening. They are used as intermediates in polymer synthe-
sis and fumigants in produce storage areas. Wastes containing phenolic
compounds are generated by the petroleum, plastics, and organic chemical
industries. Cyanide and phenolic compounds contained in wastewaters dis-
•• charged into natural water bodies may accummulate in the sediments or be
taken up by aquatic plants and animals. Materials containing cyanide or
phenolic compounds such as solid wastes from industrial and domestic
; sources, slags from mining and industrial processing, and fly ash from
- commercial operations and power generation are deposited in landfill sites.
- As a result, it is desirable to study the transport, fate, and toxicity of
' cyanide and phenolic compounds in the environment.
The goals of this research program were to develop methods for analyz-
ing solid and semisolid environmental samples for cyanide and phenolic
compounds. Preservation procedures for these samples were also investigated
to insure that the concentrations of cyanide and phenolic compounds deter-
mined in the laboratory are accurate measurements of levels present in the
environment. The program consisted of four phases:
Phase I: Selection of methodology from the literature and experi-
ence of Gulf South Research Institue and the U. S.
, Environmental Protection Agency (U.S. EPA).
Phase II: Experimentation to evaluate the methods, eliminate
nonviable alternatives, modify and optimize procedures,
and identify necessary developmental work.
Phase III: Experimentation to optimize the methods, application of
the methods to real world samples to identify potential
problems, and determination of accur&cy and precision of
the methods.
Phase IV: Statistical validation of the methods optimized in
Phase III.
I
The literature search performed in Phase I identified the gaps in
knowledge associated with preservation and analysis of solid and semisolid
environmental samples containing cyanide and phenolic compounds. The infor-
1
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nation gathered was used to develop a plan for preliminary experiments to
establish viable preservation and analysis procedures.
Separation and quantification procedures were evaluated independently
for application to solid and semisolid environmental samples. The results
of these experiments were used to identify an analytical method for cyanide
and one for phenolic compounds. During Phase III these techniques were
optimized by application to solid and semisolid environmental samples. The
methods were validated in Phase IV by the analysis of samples of ten solid
and semisolid environmental matrices. Duplicate analyses were performed on
spiked and unspiked samples of each matrix. The accuracy, precision, and
sensitivity of the methods are described.
This research effort has produced methods for the analysis of solid and
semisolid environmental samples for cyanide and total phenolic compounds.
The recommended method for determination of cyanide is distillation from an
acidified slurry of the sample followed by quantification using the pyri-
dine-barbituric acid procedure. Phenolic compounds are extracted from solid
and semisolid samples by blending with methylene chloride. The phenolic
compounds are subsequently extracted from the methylene chloride into a
sodium hydroxide solution and quantified using 4-aminoantipyrine solution.
2
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SECTION 2
RECOMMENDATIONS AND CONCLUSIONS
A review of the literature has identified the lack of information on
analysis of solid and semisolid environmental samples for cyanide and total
| phenolic compounds. The methods which were developed in this program are
! adaptations of methods for determination of cyanide and phenolic compounds
f in aqueous samples. The method for analysis of solid and semisolid environ-
! mental samples for cyanide is distillation of hydrogen cyanide from an
' acidified slurry of the sample. The distillate is bubbled through a solut-
j ion of lead acetate to remove suifur-containing compounds which would inter-
| fere in the determination procedure. The hydrogen cyanide is collected in
sodium hydroxide solution and quantified using pyridine-barbituric acid
' solution. Levels of cyanide in a sample as low as 5 pg can be detected by
1 the method. This translates to a detection limit of 0.5 (Jg/g for analysis
of a 10-g sample. The method was validated by analysis of sixty spiked and
unspiked solid and semisolid environmental samples of ten matrices. Recov-
ery of cyanide at concentrations of 2.6 to 20 |Jg/g from these samples ranged
from 69 to 112 percent, with an average of 90 + 10 percent.
The method of analysis of solid and semisolid environmental samples for
total phenolic compounds is also an adaptation of methods in the literature
< for analysis of aqueous samples. Phenolic compounds are extracted from
1 solid and semisolid samples by blending with methylene chloride. The phen-
! olic compounds are subsequently extracted from the methylene chloride into
I sodii'm hydroxide solution and quantified by the 4-aminoantipyrine procedure,
j The method was validated by analysis of sixty solid and semisolid environ-
' mental samples spiked with phenol. Recovery of phenol ranged from 22 to
| 100 percent with an average of 50 + 20 percent. Recovery appears to be
1 dependent on sample matrix, with higher recoveries being obtained from
: inorganic matrices such as sediment, fly ash, .and industrial processing
! slag. The method is sensitive to approximately 1 pg of phenol. Phenol
j concentrations as low as 0.04 pg/g can be determined in a 25-g sample.
| A round-robin study should be performed to validate the methods devel-
j oped in this program. A large number of solid and semisolid environmental
; samples should be analyzed for cyanide and total phenolic compounds.
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SECTION 3
METHOD FOR ANALYSIS OF SOLID AND SEMISOLID ENVIRONMENTAL SAMPLES FOR CYANIDE
INTRODUCTION
An extensive search of the literature was performed to find methods for
preservation and analysis of samples of all matrices containing cyanide.
Numerous methods for quantification of cyanide in aqueous samples have been
published, but limited information is available on analysis of solid or
semisolid matrices. Methods which appeared to be appropriate for applica-
tion to the analysis of solid and semisolid environmental samples were se-
lected for empirical evaluation. Selection criteria used in evaluating the
methods in the literature were accuracy and analytical range of the method,
lack of significant interferences, minimal sensitivity to changes in sample
pH or temperature, and ease of performing the determination.
Method evaluation was approached as a two-step procedure. The cyanide
was separated from the matrix and then quantified. Two separation proce-
dures, distillation and solvent extraction, were evaluated. Solid matrices
were spiked with radiolabeled cyanide and subjected to the separation proce-
dure. Liquid scintillation counting was used to determine recovery of the
radiolabeled cyanide, thus avoiding reliance on an untested quantification
method for cyanide in solid samples. The procedures evaluated for quantifi-
cation of cyanide included five colorimetric procedures, a cyanide ion-se-
lective electrode, and electron capture gas chromatography. These proced-
ures were performed on aqueous solutions containing a known concentration of
cyanide.
In the optimization phase, distillation and the pyridine-barbituric
acid colorimetric procedure were joined to form a unified method for deter-
mination of cyanide concentrations in solid and semisolid environmental
samples. A review of the literature revealed that the most significant
interference to this determination method is the positive response of vola-
tile sulfur-containing compounds. These materials were removed by bubbling
the distillate through lead acetate solution. Removal of interferences and
recovery of cyanide were monitored using the pyridine-barbituric acid color-
imetric procedure and the cyanide ion-selective electrode. Comparison of
the results of these two determination procedures, which respond differently
to the presence of sulfur-containing compounds, allowed the experimental
, conditions to be optimized to minimize interferences.
The precision, accuracy, and sensitivity of the method were determined.
Replicate samples of solid environmental matrices were analyzed for cyanide.
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These matrices were spiked with three levels of cyanide and analyzed. The
percentage of cyanide recovered and the standard deviation of the replicates
were calculated. The method was validated by anilysis of duplicate samples
of ten different matrices. These matrices were spiked at two concentrations
with cyanide and analyzed in duplicate.
PHASE I: SELECTION OF METHODOLOGY FROM THE LITERATURE
A search of the literature for information on preservation and analysis
of solid and semisolid environmental samples containing cyanide was con-
ducted in Phase I. Eighty-nine papers concerning analysis for cyanide were
retrieved using computerized and manual literature searching techniques.
While some, information has been published on determination of cyanide in
biological matrices, the majority of methods reported in the literature are
for analysis of aqueous samples. The subject of preservation of solid and
semisolid environmental samples containing cyanide has not been adequately
addressed.
The information retrieved from the literature was classified by type of
isolation or quantification procedure. The isolation procedures dr'scussed
are distillation and microdiffusion. Quantification procedures include a
variety'of colorimetric and fluorometric analyses, titrations, ion-selective
electrodes, gas chromatography, and miscellaneous instrumental techniques.
A review of the literature is presented below.
; Methods of Isolation
The term cyanide refers to aj.1 CN groups in compounds which can be
' determined as the cyanide ion, CN . There are numerous salts of cyanide,
i known as simple cyanides, which can be represented by the general formula
A(CN) where A represents an alkali or metal and x equals the valence of A
and is the number of CN groups. Cyanide salts of silver (I), mercury (I),
, and lead (II) are insoluble, whereas mercuric cyanide, HgCCN)^, is moderate-
. ly soluble in water. Cyanide metal complexes are restricted almost entirely
to the transition metals of the d block and their near neighbors zinc,
i cadmium, and mercury. The majority of cyano complexes have the general
formula
[Mn+(CN)x](x_n)", Ay(CN)x, or lM(CN)5Z]n'
where M is a metal, A is an alkali, and Z is H^O, NH^, CO, NO, H, or a
. halogen. The dissociation constants of the complexes vary greatly. The
' more refractory complexes of metals such as nickel, cobalt, and iron, re-
¦ ferred to as complex cyanides, require rigorous treatment to free the cya-
; nide ion for determination.
\ Isolation of cyanide from a matrix is generally based on the fact that
hydrogen cyanide is more volatile than other components of the sample.
Separation of total cyanide from a sample requires that all cyanide com-
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plexes be converted to the volatile form, hydrogen cyanide. Generally, only
acidic conditions are needed to degrade simple cyanides. Complex cyanides
can be broken down by heating in the presence of a catalyst or irradiating
with ultraviolet (UV) light. The addition of a catalyst and application of
heat are easily coupled with distillation and will be discussed later.
-UV irradiation has been used to decompose complex cyanides in water prior to
-'chromatographic analysis (1) or sample distillation (2-4).
Distillation—
The basic methods of isolating cyanide from a sample matrix are distil-
lation and microdiffusion. The application and rigor of these methods vary.
Distillation, commonly used to separate cyanide from aqueous samples, has
the advantage of being able to break down most complex cyanide compounds
under proper conditions. Sample pH and the presence of a catalyst are the
factors controlling the generation of hydrogen cyanide from cyanide com-
pounds. A stream of air or nitrogen is passed through the distillation
apparatus to transfer hydrogen cyanide from the sample to an absorbing
solution such as sodium hydroxide. Sulfuric acid is commonly used to adjust
.the pH of the sample (A—9). Tartaric acid (7,10), hydrochloric acid (11),
hypophosphoric acid (3), phosphoric mercaptoacetic acid (12), and sodium
bicarbonate (11) have also been used to lower sample pH. The recovery of
simple and complex cyanide from body fluids by distilling the sample in the
presence of hydrochloric acid, sodium bicarbonate, and pH 6.5 buffer salt
was studied (11). Recovery of cyanide was found to increase with a decrease
in pH.
Catalysts such as magnesium chloride (7,13-15), mercuric chloride
(7,14,15), and cuprous chloride (9,13,16) have been added to the sample to
increase the recovery of cyanide from refractory compounds. The recovery of
simple and complex cyanides from aqueous samples distilled in the presence
| of sulfuric acid and the catalysts magnesium chloride and mercuric chloride
; (Serfass distillation), and tartaric acid with no catalysts was studied (7).
j All simple cyanides and complex cyanides ot zinc, cadmium, nickel, and
' silver are easily distilled by either procedure. Ferro- and ferricyanides
can also be recovered quantitatively by either method. Recovery of complex
• copper cyanides was high only with Serfass distillation, while recovery of
• cobalticyanides was only 20 percent with this method and very low with
; tartaric acid distillation (7). Formation of mercuric cyanide which is not
recovered by distillation if the chloride concentration is insufficient can
' be prevented by using a higher ratio of magnesium chloride to mercuric
i chloride (14). Over 99 percent of the ferrocyanide in water can be recover-
| ed by distillation in the presence of cuprous chloride and sulfuric and
' hydrochloric acids (16). The method recommended by the American Public
: Health Association (APHA) (13) calls for addition of sulfuric acid and
< cuprous chloride or magnesium chloride to the sample prior to distillation,
; while the method recommended by the U.S. EPA (9) utilizes magnesium chloride
j and sulfuric acid.
! The apparatus used and length of time allowed for distillation of
i cyanide vary. The most commonly used apparatus (3,7,9,11,13,14) for frac-
6
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tional distillation was devised by Serfass (15) which consists of a distil-
lation flask equipped with an Allihn condenser attached to a gas washer and
a vacuum supply (Figure 1). Steam distillation has been used to separate
cyanide from blood and tissue (17). Simple cyanide can be separated from
ferrocyanide by distillation under reduced pressure (18). The distillation
process has been automated (2), and a microstill has been used for isolating
"hydrogen cyanide from insects (10).
Aeration has been used as a method for separation of hydrogen cyanide
from biological (19-24) and water samples (25). The sample is acidified,
and held at room temperature or warmed. A stream of air or nitrogen is
passed through or over the sample to sweep the hydrogen cyanide into an
absorbing solution or onto a packed column. The major factors affecting the
efficiency of isolating hydrogen cyanide from samples by aeration have been
found to be sample pH and temperature (21).
Microdiffusion—
A frequently used method of isolating cyanide from biological samples
is microdiffusion (26-32). A sample of blood, urine, gastric contents, or
homogenized tissue and an aliquot of sulfuric acid are placed in the outer
chamber of a Conway microdiffusion cell. Sodium hydroxide solution is
placed in the inner chamber,.and the cyanide is allowed to diffuse from the
sample into the sodium hydroxide solution. The Cavett blood-alcohol flask
has been modified for uje as a diffusion cell (31). The microdiffusion
technique isolates only volatile components from the sample; therefore, it
can not be used to determine total cyanide without sample pretreatment. The
volume of sample which can be contained in the microdiffusion cell is small
(less than 5 ml); thus, the detectable concentration in the sample is se-
verely limited by sample size.
Methods of Quantification
The majority of methods for quantification of cyanide are based on the
reaction of cyanide ions with a colorimetric reagent. In general, these
spectrophotometric methods are accurate and sensititive. Fluorometric
methods have been developed which have sensitivities companble or superior
to spectrophotometric methods, but they require more sophisticated instru-
mentation. Titration methods, while accurate, are not sensitive enough to
determine traces of cyanide in environmental samples. Satisfactory instru-
mental methods using ion-selective electrodes, gas chromatography, coulo-
metry, and atomic absorption spectrophotometry have been developed in recent
years.
Spectrophotometric Methods—
Konig reaction—A basic method for the ^determination of cyanide in
aqueous (33-36) and biological samples (11,23,26) is based on a Konig reac-
tion (37). A common method (37) utilizes Konig synthesis of a pyridine dye
in which cyanogen bromide or cyanogen chloride reacts with pyridine and an
( aromatic amine to form a dye. The cyanide is brominated, excess bromine is
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J
Figure 1. Apparatus for distillation of cyanide.
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removed with arsenious acid, and the cyanogen bromide reacts with a mixed
pyridine-benzidine reagent. The absorbance of the dye is proportional to
the concentration of cyanide in the sample. Concentrations of cyanide as
low as 0.1 mg/L can be determined with an error of ± 2 percent (38). This
method has been used to determine cyanide in blood, urine, and pancreatic
juices deproteinized with trichloroacetic acid (22,38) or distilled with
acid. (11). A comparison of seven amines with pyridine and forty-six aro-
matic amines with benzidine was made in a search for noncarcinogenic rea-
gents which are more stable and have a greater molecular extinction coeffic-
ient than pyridine and benzidine (33,34). By substituting p-phenylene-
diamine for benzidine and slightly altering the procedure, concentrations of
cyanide ranging from 0.005-100 mg/L can be determined (39). In recent years
this method has been used for analysis of biological fluids (23,26,40) and
adapted for automated analysis of water and wastewater (35,36).
The Konig synthesis using pyridine and benzidine or pyridine and £-phe-
nylenediamine is sensitive to thiosulfate and sulfide as well as cyanide.
Interference from thiocyanate, turbidity, and color can be eliminated by
distillation of the sample, and sulfide can be removed by precipitation with
cadmium. The intensity of the dye is sensitive to the pH and temperature of
the test solution and the time allowed for development of the dye. These
factors can be controlled easily enough to render the method viable.
The intermediate cyanogen chloride can be replaced with the less vola-
tile cyanogen bromide, and barbituric acid can be used as the condensing
agent for the reaction with pyridine (41). The stability of the dye formed
using cyanogen bromide with pyridine and barbituric acid is superior to that
of the dye formed in the cyanogen chloride-barbituric acid method or the
benzidine or pyrazolone methods. Recovery of 96-102 percent at cyanide
levels of 0.1-2.0 pg in plasma, urine, and tissue samples using cyanogen
chloride with pyridine and barbituric acid has been reported (28). The
American Public Health Association (13) and U.S. EPA (9) also advocate the
cyanogen chloride-pyridine-barbituric acid method for determination of cya-
nide in water samples.
A method (42) for determination of cyanide in water is recommended by
the U.S. EPA (9) as an alternative to the pyridine-barbituric acid method
and has been used by many for the determination of cyanide in wastewater
(2,18,43). Chloramine-T is used to convert cyanide to cyanogen chloride.
The cyanogen chloride reacts with a solution of pyridine containing 0.1 per-
cent bis-pyrazolone and l-phenyl-3-methyl-5-pyrazolone to form a dye which
is stable for 30 minutes at 25°C. The absorbance of the dye measured at
630 nra is proportional to the concentration of cyanide in the sample. The
advantages of this method are that as little as 0.2 pg of cyanide can be
determined with an accuracy of 99 ± 4 percent, and that there is no change
in intensity of the dye with pH changes in the range 2.8-9.0. High concen-
trations of reducing agents may interfere with the determination (42). This
method has been adapted to the analysis of biological samples including
blood, urine, tissue,- and gastric contents (3,14,19,21,28,31). The pyri-
dine-pyrazolone dye can be extracted with butyl alcohol to increase the
.9.
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sensitivity of the method (7,44). The color reaction is generally repro-,
ducible over a broad range, but a significant buffer capacity of a sample
will decrease the effective pH range (7,44).
(
Metallic reactions—One of the earliest methods for determination of
.cyanide involving formation of a metal complex was based on the formation of
Prussian blue, Fe^lFe(CN),]^, by the reaction of cyanide with ferrous sul-
fate (10,45). This method is sensitive (0.01 mg hydrogen cyanide), but has
many shortcomings. The age of the ferrous sulfate solution causes the color
to vary significantly. The presence of air during the reaction period
influences the size of the colloidal particles and thus the photometer
readings. Improved methods of analysis based on the reaction of cyanide
with 1,10-phenanthroline were reported (46-48). A method based upon for-
mation and extraction of thr neutral dicyano-bis(l,10-phenanthroline) iron
(II) complex produced by the exchange reaction between tris(l,10-phenan-
throline) iron (II) and cyanide ions uas developed (46). The color produced
is stable for up to an hour. Disadvantages of the method include inter-
ference by periodate, cobalt, copper, nickel, and ferrius ions; incomplete
recovery of the dicyano-bis(l,10-phenanthroline) iron (II) complex by ex-
traction; and extreme sensitivity to variation of pH (46).
Another procedure is based on the reaction of tris(l,10-phenanthroline)
iron (II) triiodide adsorbed to porcelain chips with cyanide to release the
red complex cation (47). Absorbance at 514 nm by the red complex is linear
up to 8 mg/L. None of several ions added to the sample interfered with the
determination.
An indirect spectrophotometric method is based on the fact that cyanide
prevents formation of the strongly absorbing ternary complex of silver (I),
1,10-phenanthroline and bromopyrogallol red in nearly neutral aqueous solu-
tion (48). A plot of absorbance versus concentration is linear over the
range 0.26-2.6 mg/L of cyanide. Several anions and cations interfere with
the determination, but most of these can be eliminated by precipitation or
masked with EDTA.
The application of mercury chloranilates > for the spectrophotometric
determination of cyanide was evaluated (49). The reaction of cyanide with
mercury chloranilate forms soluble, undissociated mercuric cyanide. Cyanide
concentration can be determined over the range of 0.4-4.0 mg/L by measuring
the absorbance of the solution at 330 nm. The method is hindered by the
significant interference of sulfite and sulfide.
A procedure for quantification of cyanide using ammoniacal nickel
chloride was developed (5). Hydrogen cyanide is distilled from the sample
and collected in a solution of ammoniacal nickel chloride. The cyanide
reacts immediately to form an ionic complex which can be quantified by
ultraviolet spectrophotometry. Hydrogen sulfide and sulfite must be removed
prior to analysis to prevent interferences. The method follows Beer's law
over the range of 0.1 to 4.0 mg/L of cyanide.
-------
Organic disulfide reactions—The reaction of cyanide with four disul-
fides to displace a thiol anion was evaluated as a spectrophotometry method
for determination of cyanide (50). N,N-dimethylformamide is added to the
test solution to increase the rate of reaction and improve sensitivity (51).
Absorbance of the thiol anion is measured and related to the cyanide concen-
tration over the range of 2.6 to 26 mg/L. Ions of cadmium, copper, iron,
mercury, nickel, lead, and zinc interfere at concep.tidtions of 20 mg/L with
the determination of 1 mg/L cyanide. EDTA can be added to eliminate inter-
ference by all of the cations except copper and mercury. Interference by
mercaptans, sulfide, and sulfite can be minimized by adjusting the pH.
Fluorometric Methods—
Demasking reactions—Demasking reactions which form fluorescent com-
pounds have been used for determination of cyanide (52-55). A qualitative
test for cyanide based on the demasking of dimethylgloxime by the reaction
of cyanide with palladium (II) dimethylgloxime was developed (53). Nickel
in the solution reacts with the dimethylgloxime to form the red nickel
dimethylgloxime. A fluorescent spot test for detecting cyanide in vapor
based on the demasking of oxine from copper (II) oxinate which permits
formation of a fluorescent aluminum oxinate compound was proposed (52). A
quantitative method for analysis of trace levels of cyanide based on the
demasking of 8-hydroxy-5-quinolinesulfonic acid by cyanide from the non-
fluorescent potassium bis(5-sulfoxino) palladium (II) was reported (54).
The liberated 8-hydroxy-5-quinolinesulfonic acid coordinates with magnesium
ions to form a fluorescent chelate, the inten&ity of which is proportional
to the amount of cyanide present. This method has been adapted to the
automated determination of cyanide in blood (24). It was found that the
demasked 8-hydroxy-7-iodo-5-quinolinesulfonic acid) formed by reaction of
cyanide with potassium bis-(7-iodo-5-sulfoxino) palladium (II) forms a
stable blue-green color upon addition of a ferric salt (54). The fluoro-
metric method is approximately two orders of magnitude more sensitive than
the colorimetric method. Achievement of accurate and precise results with
the fluorometric method requires careful control of pH and reaction time.
Both methods respond to sulfide, and the colorimetric reaction responds to
thiocyanate, thiols, and certain disulfides as well.
The reaction of cyanide with the nonfluorescent bis-palladiura-bis-
(2,3-diaminonaphthalene-selenium)tetrachloride (DANSe) to demask the fluo-
rescent DANSe was proposed as a method for quantifying cyanide (55). The
method can detect concentrations of cyanide as low as 0.6 mg/L.
Other reactions--Fluorescent compounds can be produced by the addition
of cyanide to the appropriate anion (56,57), cleavage of certain aromatic
rings by cyanide (58), and formation of a specific compound by reaction with
cyanide (29). Addition of as little as 0.5 pg of cyanide to quinone mon-
oxime benzene sulfonate ester produces a highly fluorescent compound (57).
The fluorescence produced by the reaction of £-benzoquiDone with cyanide is
proportional to the cyanide concentration over the range 0.2 to 50 pg (56).
The fluorescence is most intense when dimethylsulfoxide is used as the
solvent. No ions interfere in concentrations up to 0.1M.
-------
A method for determination of cyanide based on cleavage of tho pyridine
ring of nicotinamide by cyanogen chloride was developed (58). The fluo-
rescent intensity of the cleaved ring is dependent on the pH of the solution
and time allowed for reaction, but the method is suitable for determination
of 0.3 to 6 rag/L of cyanide.
-Catalytic reactions can be used for quantification of cyanide. The
oxidation of pyridoxal with oxygen by the catalytic action of cyanide to the
fluorescent 4-pyridoxalactone was the basis of a method (59). The finding
that the reductant cyanohydrin, formed by reac.jkion of cyanide with j>-nitro-
benzaldehyde, is capable of reducing a variety of compounds to form a highly
colored product has been the basis of several methods (60). The catalytic
reaction of cyanohydrin with o-dinitrobenzene to give a blue compound can be
used to quantify the cyanide by measuring the change in absorbance over
time (60). Cyanide in tissues has been determined by the reaction of cyano-
hydrin with £-benzoquinone in dimethyl sulfoxide to form a fluorescent
product (29). The relationship of fluorescence intensity and cyanide con-
centration is not linear at low concentrations; thus, the sensititivity is
.limited to 1 mg/L.
While fluorometric methods are often more sensitive than colorimetric
methods, they have several disadvantages which tend to make them less appli-
cable than other methods for routine analysis of environmental samples. As
with colorimetric methods, most fluorometric methods are very sensitive to
the pH of the sample solution. Also critical to obtaining reproducible data
is the requirement that the fluorescence be measured after a precise inter-
val of time. Frequently the compounds used in fluorescent reactions are
unavailable commercially and must be synthesized or purified in the labora-
tory.
Titrimetric Methods—
The standard method for quantifying cyanide at concentrations greater
than 1 mg/L is titration with silver nitrate solution (9,13). The endpoint
is detected by the silver-sensitive indicator £-dimethylaminobenzalrhodanine
(9,13,61) or by a silver ion-selective electrode (62). Others have titrated
the sample with Hg-EDTA using mercury (II) cresol-phthaleincomplexone-ioaide
as the endpoint indicator (63) and with chloramine-T, dichloramine-T, and
lead tetraacetate using a platinum indicator electrode and a reference
calomel electrode to detect the endpoint (64). Cyanide in a solution can be
determined indirectly by oxidation with chloramine-T or dichloramine-T. An
excess of potassium iodide is added to the solution and titrated with sodium
thiosulfate (65). Another method is to oxidizo the cyanide with bromine,
remove excess bromine with phenol, and determine the cyanogen bromide iodo-
metrically (12). The performance of a cation-selective glass electrode and
a silver metal electrode were compared for detecting the endpoint in argen-
tometric determination of cyanide (66). It was found that the glass elec-
trode was superior for titration purposes.
Ion-Selective Electrodes—
A cyanide or silver ion-selective electrode can be used to quantify the
^concentration of free cyanide ion in a solution (67-70). The activity of
_
12
-------
-the cyanide ion is determined by the electrode; therefore, the ionic
strength of samples must be controlled (68,70). The easiest way to accom-
plish this is by diluting samples and standards 1:1 with 2M sodium hydroxide
which serves to swamp out variations in the original total ionic strength
(6,70). Another approach to determining the cyanide concentration with an
...ion-selective electrode is the method of known additions. A known amount of
"cyanide, is added to a known volume of sample, and the change in electrode
potential is used to calculate the cyanide concentration (68). Principal
interferences are the sulfide and iodide ions (70). Reported limits of
detection range from 0.02'mg/L (68) to 0.3 mg/L (70). The cyanide ion-
selective electrode has been used to quantify cyanide in water (71), indus-
trial effluents (68,70,72,73), plant homogenates (73), and fruit and plant
distillates (75). A comparison of the determination of cyanide in steelJ
works effluents by the cyanide ion-selective electrode, argentometric ti-
tration, pyridine-pyrazolone colorimetric method, and pyridine-barbituric
acid colorimetric method was made (72). The electrode method and argento-
metric titration gave comparable results for cyanide concentrations greater
than 1 mg/L. At lower levels, the pyridine-pyrazolone method tended to give
higher results than the other methods. Quantification of cyanide ion by the
electrode is a quick, simple method when the concentration is in the optimal
range and no interferences are present.
Gas Chromatographic Methods—
Gas chromatography (GC) can be used to quantify cyanide in gases (76),
water (1,25,77), blood and urine (27), and alcoholic beverages (6). Celite
or Chromosorb coated with 20 percent polyethylene glycol 1500 is satis-
factory for separating hydrogen cyanide from water vapor and permanent gases
(76). Hydrogen cyanide ranging in concentration from 0.026 to 13 mg/L can
be purged from solution and concentrated on a column packed with 19.A per-
cent dinonyl phthalate on 40/60 mesh firebrick (25). Compounds which are
not separated from the hydrogen cyanide and, thus, interfere are trimethyl-
amine, methylamine, diethylamine, methanol, pentanes, and pentenes (25).
Another approach is bromination of the cyanide followed Dy injection of the
aqueous solution (1,77) or a diisopropyl ether extract (6) onto a Porapack Q
column. The minimum cyanide concentration in the sample which can be
detected by the electron capture detector was reported to be 10 lig/L for
direct aqueous injection (1,77) and 1 |Jg/L for the ether extraction proce-
dure (6).
Miscellaneous Instrumental Methods— ,
Instrumental methods of analysis for cyanide which have received limit-
ed use include coulometry (78), polarography (79), and atomic absorption
spectrophotometry (AAS) (80). Cyanide at levels ranging from 13 to 260 [lg/
sample has been determined by coulometric titration with electrogenerated
hypobromite (78). The technique is accurate and rapid but very sensitive to
slight_^H changes. Determination of cyanide ion at levels as low as
2 x 10 Mg/mL was achieved using a polarographic method (79). The
response of a rapidly rotating gold electrode to cyanide ion in lithium
hydroxide was used to quantify the cyanide in the aqueous sample. Cyanide
can be determined indirectly by AAS. The cyanide is allowed to react with a
13
-------
metal, and the resulting cyanide-metal complex is quantified by AAS.
Examples of this method include precipitation of cyanide with silver and
formation of a dicyano-bis(l,10-phenanthroline) iron (II) complex (80)
Selection of Methods
.Methods which appeared to be suitable for adaptation to analysis o
solid and semisolid environmental samples for cyanide were selected for
empirical evaluation in Phase II. Distillation and microdiffusion can be
used to separate free cyanide from a liquid matrix. Distillation has an
advantage over diffusion in that heat and catalysts can be applied to aid
the breakdown of complex cyanides to permit determination of total cyanide
in the sample. Efficient removal of cyanide from relatively large volumes
of sample can be achieved with distillation. For these reasons, distilla-
tion was selected as one of the separation procedures for evaluation for
analysis of trolid samples. The other separation procedure chosen was
ion-pair extraction. While no references were found for extraction of
cyanide as an i>n pair, the technique warranted investigation (81).
Selection criteria used in evaluating quantification methods in the
literature were analytical range and accuracy of the method, ease of deter-
mination, lack of significant interferences, and minimal variation of ana-
lytical response as a function of sample pH or temperature. The quanti-
fication methods which were evaluated are listed below.
1. The colorimetric method based on Konig synthesis of a dye by the
reaction of cyanide with chloramine-T to form cyanogen chloride
which subsequently reacts with pyridine-barbituric acid reagent
(9).
2. Colorimetric determination based upon formation and extraction of
the neutral dicyano-bis(l,10-phenanthroline) iron (II) complex
produced through the exchange reaction between tris(l,10-phenan-
throline) iron (II) and cyanide ions (46).
3. Ultraviolet spectrophotometric determination based upon the ab-
sorbance of the chloranilate ion formed by reaction of mercuric
chloranilate with cyanide (49).
4. Ultraviolet spectrophotometric quantification of the tetracyano-
nickelate (II) anion complex formed by reaction of hydrogen cya-
nide with amraoniacal nickel chloride (5).
5. Spectrophotometric determination of the thiol anion which is
displaced by reaction of N,N-dimethylformamide with cyanide ions
(51).
6. Direct determination of cyanide ion activity using an ion selec-
tive electrode.
\
.14
-------
I
7. GC determination of cyanogen bromide formed by bromination of
cyanide in the sample (77).
PHASE II: EVALUATION OF THE METHODS
The goals of this phase were to eliminate nonviable methods and iden-
tify necessary modifications of promising procedures. Analysis of a solid
ar semisolid environmental sample for cyanide was approached as a two-step
procedure: separation of cyanide from the sample matrix and quantification
of the cyanide. Experiments w» re performed to evaluate two separation and
seven determination procedures. Distillation and solvent extraction as an
ion pair complex were the separation procedures studied. Of the seven
determination procedures, five were spectrophotometry procedures based on
measurement of the absorbance of a compound formed by reaction of a reagent
with cyanide. The other procedures were quantification using a cyanide
ion-selective electrode and a gas chromatograph equipped with an electron
capture detector. 1
Three solid or semisolid matrices were used for the evaluation of sep-
aration procedures. The matrices were selected to cover the range of char-
acteristics encountered in solid and semisolid environmental samples. Lake
sediment was used as a representative inorganic matrix. Fish tissue was
employed as a matrix of high organic content. Organic domestic sludge from
a sewage treatment plant drying pond, the third matrix, was characterized as
being an organic material containing significant levels of trace metals.!
In all experiments evaluating separation proc^ures, samples were
spiked with a sodiuij^ hydroxide solution containing C-labeled potassium
cyanide. The use of C-labeled cyanide permitted determination of recovery
^hieved by separation procedures by liquid scintillation counting of the
C. Using this technique avoided the necessity of having to rely on deter-
mination procedures designed for analysis of water samples which had not
been ^ilidated for solid and semisolid samples. The assumption was made
that C-labeled cyanide responds exactly as unlabeled cyanide to separation
and determination procedures. The comparability of liquid scintillation
counting and a conventional analytical procedure was demonstrated by using
liquid scintillation counting and a cyanide ion-selective electrode to
quantify cyanide in 2 sets of 10 replicate samples. The average concen-
trations of cyanide measured in the first set were 0.75 mg/L ± 0.04 and
0.76 mg/L ± 0.12 with liquid scintillation counting and the ion-selective
electrode, respectively. Analysis of the second set yielded 0.77
mg/L ± 0.04 and 0.82 mg/L ±0.11 using liquid scintillation counting and the
ion-selective electrode, respectively.
Separation and determination procedures were evaluated by performing
the procedures on seven replicate samples. The percent recovery and stan-
dard deviation of the replicates were calculated using the following
equations:
.15 _
-------
Percent recovery =
100 (0.)
T.
1
where 0.
l
observed value
T. , . true value
l;
n
where n = number of samples
= percent recovery of sample i
Standard deviation = I—=-
1/ n-1
n 2 . n 2.
I P. - I P. / n
i=l 1 li=l
Modifications were often made of procedures which were determined to be
viable. Procedures and modifications of procedures which were judged to be
unsatisfactory by initial experiments were not always tested by performing
the procedure on seven replicates.
Procedures for Isolation
Isolation procedures were tested on samples of fish tissue, sediment,
and sludge. Large quantities of each matrix were obtained and processed as
described below to provide material for all evaluation experiments. Fish
tissue was prepared for analysis by chopping the edible portions of fresh
fish into small pieces and homogenizing in a blender. The tissue was stored
in a freezer until needed. Lake sediment was dried at 103°C and then clean-
ed of large pieces of debris such as sticks, leaves, and shells. Sludge was
dried at 103°C, pulverized with a mortar and pestle to generate a powdery
matrix, and stored at room temperature.
A procedure was developed for spiking samples of the solid matrix prior
to analysis. Solid and semisolid samples were moistened with sodium hy-
droxide solution and then spiked with a small volume of sodium hydroxide
solution containing radiolabeled potassium cyanide. Sludge and sediment
samples were held at room temperature for approximately 16 hours to allow
time for adsorption of the cyanide to the solid particles. Fish tissue
samples were held for 16 hours at 4-6°C to prevent decomposition of the
tissue. Control samples were prepared with each set of solid samples by
adding aliquots of the sodium hydroxide solution and cyanide spiking so-
lution to an appropriate volume of water. Controls were treated by the same
procedure as the solid samples and were identical to the solid samples
except that they contained no solid matrix. '
16
-------
^Liquid scintillation counting was used to determine tlj^ concentration
of C in all solutions. Aliquots of solutions containing C were counted
at various steps of the procedures to determine recovery of cyanide. The
concentration of radiolabeled cyanide was determined by placing 0.5 mL or
1.0 mL of solution in a glass vial containing 10 mL of liquid scintillation
cocktail and counting with a liquid scintillation counter. Three replicate
-aliquots of each sample were always counted, and the average was reported.
Blanks were prepared by taking an un^iked sample or control through a
separation procedure. The levels of C found in the blanks were never
above expected background.
Distillation—
Before work with solid samples was initiated, experiments were per-
formed to verify that the distillation apparatus and the experimental tech-
niques used would give high recovery of cyanide distilled from water. The
distillation apparatus consisted of a one-liter round bottom distillation
flask equipped with a condenser, air inlet tube, and absorber tube. A
vacuum system was used to pull air into the flask through the air inlet tube
and out through the absorber tube. It was determined that the air flow rate
is a significant factor affecting the recovery of cyanide. Recoveries above
80 percent were obtained when the air flow rate was great enough to produce
a layer of foam 2 cm high on the top of the sodium hydroxide solution in the
absorber tube.
The effect of holding time between spiking and distilling the samples
on the recovery of cyanide was briefly investigated. The relationship of
recovery of cyanide to the length of holding time between spiking and dis-
tilling was studied by spiking four sediment samples and four controls
containing no sediment as described above. Distillation of two samples and
two controls was initiated 0.5 hour after completion of spiking. The re-
mainder of the samples was held 16 hours and then distilled. The purpose
of this experiment was to determine if the percent accounted for was lower
for samples held overnight because of loss of cyanide from the sample and
flask. As shown in Table 1, the percent accounted for is similar for sam-
ples and controls held for 0.5 hour and 16 hours. Recovery of cyanide from
samples held for 16 hours may be lower than that of samples held for
0.5 hour because of increased adsorption of cyanide to the solid matrix, but
cyanide does not appear to be lost from the system.
Two basic procedures were evaluated for separating cyanide from solid
and semisolid environmental samples. The procedures were distillation and
ion pair solvent extraction. Several variations of distillation were tried.
These involved stirring the sample with a solution of sodium hydroxide prior
to distillation, stirring the sample with sulfuric acid and reagents during
distillation, or substituting mercuric or cuprous chloride for the magnesium
chloride refluxing reagent.
-------
TABLE 1. COMPARISON OF RECOVERY OF CYANIDE FROM SEDIMENT HELD
0.5 HR AND 16 HR BEFORE DISTILLING
Holding time
Percent
Total
Number
between spiking
recovered
Percent
percent
of
and distilling
by 1 hr
remaining
accounted
Sample
replicates
(hr)
distillation
in flask
for
Sediment
2
0.5
86 + 1
16 + 0
102 + 1
Sediment
2
16
78 + 0
24 + 0
102 + 0
Control
2
0.5
86 + 8
14 + 0
100 + 8
Control
2
16
94 + 2
16 + 0
110 + 2
Experiment's designed to assess recovery of cyanide from solid and
semisolid samples by distillation were performed. Ten-gram portions of
sludge were spiked with 100 |jg of radiolabeled cyanide as potassium cyanide
in a sodium hydroxide solution. Ten-gram samples of fish tissue were
spiked with 50 |jg of radiolabeled cyanide. Controls were prepared in the
same manner but contained no sludge or fish tissue. Immediately prior to
distillation, 250 mL of distilled water, 10 mL of 5M magnesium chloride
solution, and 12 mL of concentrated sulfuric acid were added to the sludge
or fish tissue in the distillation flask. The flask was attached to an
absorber tube filled with 50 mL of 1.25M sodium hydroxide solution. Air was
bubbled through the distillation system at a rate great enough to maintain
about 2 cm of foam on the surface of the sodium hydroxide solution in the
absorber tube. It was desirable to know the time period required for recov-
ery of all of the cyanide from the samples. A set of samples was distilled
for 30 minutes, fresh sodium hydroxide solution was placed in the absorber
tube, and the samples were disLilled for a second 30-minute period. The
sodium hydroxide solution was again replaced, and the samples were distilled
for a second hour. The average percentage of cyanide recovered from each
matrix for each distillation period is shown in Table 2. Recovery after 1
hour of distillation ranged from 35 percent for fish tissue and 39 percent
for sludge to 90 percent for controls. An additional 9 percent of the
cyanide was recovered from samples of sludge and fish tissue during the
second hour of distillation.
The distillation procedure was modified to increase the recovery of
total cyanide from solid samples. Five- or ten-gram portions of fish
tissue, sediment, and sludge were spiked with 50 to 200 |Jg of radiolabeled
cyanide as potassium cyanide. The samples were held overnight and then
stirred with 100 mL of 2M or 4M sodium hydroxide solution for 1 hour. The
mixture was transferred to a distillation flask, and the volume was brought
up to 500 mL with distilled water. Twenty-five or fifty milliliters of
concentrated sulfuric acid and 20 mL of 5M magnesium chloride solution were
added to the distillation flask. The mixture was boiled gently, and a flow
of air was bubbled through the system to carry hydrogen cyanide from the
.mixture to an absorber tube containing 50 mL of 1.25M sodium hydroxide
-------
-solution. The samples were distilled for 1 hour, the sodium hydroxide
solution in the absorber tube was replaced with fresh solution, and the
distillations were continued for a second hour. The average percentages of
cyanide recovered by the sodium hydroxide stirring and distillation proce-
dure are shown in Table 2. Recovery tended to be about the same or
.slightly lower than recovery achieved by distillation alone. Recovery of
'-cyanide from fish tissue was 28 percent for 1 hour of distillation and
36 percent for 2 hours. Thirty-eight and 43 percent were recovered from
sludge by 1 and 2 hours of distillation, respectively. The recovery of
cyanide from sediment was 64 percent after 1 hour with no increase after
2 hours of distillation.
TABLE 2. PERCENTAGES AND STANDARD DEVIATIONS OF CYANIDE RECOVERED
FROM THREE MATRICES BY DISTILLATION PROCEDURES
Matrix
Duration
of
procedure (hr)
Distillation
Distillation
preceded by
stirring with
NaOH solution
Distillation
with
stirring
Fish
tissue
35 ±
44 ±
4
4
28+4
36 ± 4
Sediment
1
.2
64 ± 15
63 ± 16
69 ± 15
71 ± 10
Sludge
0.5
1
2
30 + 1
39 + 6
38 ± 6
43 ± 4
43 ±
52 ±
Controls
(water)
0.5
1
2
76
90 ± 2
93
85 ± 7
84 ± 20
103 ± 4
Another set of experiments was performed to determine whether higher
recoveries could be obtained by stirriag the slurried sample during dis-
tillation. Stirring prevents sampl» particles from settling to the bottom
of the distillation flask and reduces bumping of the flask. Ten-gram por-
tions of dry sediment were spiked with 47 |Jg of radiolabeled cyanide as
potassium cyanide in a sodium hydroxide solution. The sludge samples were
prepared in a similar manner, but 5-g portions of dry sludge were spiked
with 47 pg of radiolabeled cyanide. Controls were prepared for the sludge
and sediment samples by adding the sodium hydroxide solution and radio-
labeled potassium cyanide to an empty distillation flask. Immediately prior
to distillation, 500 mL of distilled water, 25 mL of concentrated sulfuric
acid, and 20 mL of 2.5M magnesium chloride solution were added to the sam-
ples. The flasks were connected to absorber tubes filled with 50 mL of
19
-------
1-.25M sodium hydroxide solution, and the samples were stirred with a mag-
netic stirring bar and distilled for 1 or 2 hours.
Recovery of cyanide from the sediment and sludge samples was slightlj
higher than that measured in previous experiments as shown in Table 2.
.Seventy-one percent of the cyanide was recovered from the sediment an<
;52 percent from the sludge by 2 hours of stirring and distilling. The
greatest advantage of this procedure is that stirring tends to decrease
bumping of the distillation flasks- to make distillation of a larger size
sample feasible.
A series of experiments was performed to determine whether substituting
solutions of cuprous chloride or mercuric chloride for the magnesium chlo-
ride refluxing reagent would improve the recovery of cyanide. Five grams of
sludge or 10 g of sediment or tissue were spiked with 50 pg of radiolabeled
cyanide as potassium cyanide in a sodium hydroxide solution. A 500-mL
volume of distilled water; a specific volume of magnesium chloride, cuprous
chloride, or mercuric choloride; and 25 ml of concentrated sulfuric acid
were added to the samples. The flask was connected to an absorber tube
filled with 50 roL of a 1.25M sodium hydroxide solution, and samples were
distilled for 1 hour unless otherwise noted.
Details of the experiments and the recovery of cyanide as determined by
liquid scintillation couating are presented in Table 3. Comparison of the
average percentage of cyanide recovered using the magnesium chloride reagent
with that using cuprous chloride or mercuric chloride reagents showed no
significant difference. Distillation for a second hour increased the recov-
ery by an average of 4 percent for sediment, 6 percent for sludge, and
17 percent for fish tissue.
Solvent Extraction—
In addition to distillation, solvent extraction was evaluated as a
procedure for separating cyanide from solid matrices. Preliminary experi-
ments to evaluate the procedur^were performed on samples of dilute sodium.
hydroxide solution containing C-labeled and unlabeled potassium cyanide.
Ten-milliliter aliquots of solution containing 5 )Jg of cyanide and 1-mL
aliquots of 0.001M ammonium hydroxide were place in test tubes. The ammo-
nium-cyanide ion pair was extracted from aqueous solution with hexane or
methylene chloride. Recoveries of cyanide were determined by liquid scin-
tillation counting of the extraction solvent and the samples before and after
extraction. Less than 1 percent of the cyanide was extracted by either
solvent. No further work was performed with this procedure. Initial exper-
iments showed it to be unsatisfactory, and problems with extraction of
cyanide as an ion-pair from solid samples were anticipated.
Procedures for Quantification
Seven analytical procedures for quantification of cyanide were selected
from the literature for evaluation. Criteria used for selection were ana-
lytical range of the procedure, lack of significant interferences, and
minimal variation of analytical response as a function of pH, temperature,
20
-------
TABLE 3. RECOVERY OF CYANIDE FROM FISH TISSUE, SEDIMENT AND SLUDGE BY DISTILLATION
WITH SEVERAL REFLUXING REAGENTS
Average percent Average percent recovered +
Number recovered ± standard deviation by 1 br
of standard deviation of distillation using
" Matrix "" replicates Volume of reagent 1 hr 2 hr only MgC^
Fish tissue
3
40
ml
of
2.5M MgCl2 +
28 +
4
45
+
1
35 +
4
10
ml
of
0.25M HgCl2
-
1
Sediment
3
10
ml
of
0.2M CuCl
61 +
10
j
65
+
9
65 +
17*
Sludge
-¦3-~
40
ml
of
2.5M MgCl2 +"~
34 +
3
¦ 40
+
2
i
37 .+
8 -- -
10
ml
of
0.25M HgCl2
^Samples werestirred with sodium hydroxide solution prior to distillation.
-------
and .reaction time. Evaluation of determination procedures was performed
using solutions of potassium cyanide prepared in dilute sodium hydroxide
solution. Solutions were prepared fresh daily by dilution from a stock
solution of 1000 mg/L cyanide. The stock cyanide solution was standardized
by titration with a standard solution of silver nitrate. Evaluation of the
procedures was based on precision and accuracy data generated by analysis of
aqueous solutions of potassium cyanide. Seven replicate samples of concen-
tration approximately ttn times the limit of detection were analyzed by each
procedure. The average percent recovery and standard deviation were calcu-
lated for all of the procedures except quantification by GC and are present-
ed in Table 4. Complete information on reagents and steps of the proce-
dures, which are listed on pages 14 and 15, are included in Appendix A
The GC procedure was not thoroughly evaluated, because of the poor re-
covery of the ion-pair extraction procedure prescribed above and the numer-
ous problems encountered with the GC procedure itself. However, preliminary
evaluation was made of the GC quantification procedures which consists of
2 steps, formation of cyanogen bromide and analysis by GC.
Cyanide in samples which were to be analyzed by GC was reacted with
bromine to form cyanogen bromide which was then extracted into an organic
solvent. The following procedure for bromination (77) and extraction of
cyanide in aqueous samples was evaluated. Replicate 50-mL aliquots of
0.1M potassium hydroxide solution containing 5 |Jg of radiolabeled cyanide as
potassium cyanide were treated with bromine and extracted by the following
procedure. The pH of the solution was adjusted to 4 or 5 with 20 percent
phosphoric acid. Cyanogen bromide was formed by adding saturated bromine
water to the solutions drop-by-drop until a yellow color persisted. An
excess of 5 drops was added; the sample was shaken to mix and held for
5 minutes. A sufficient volume of a 5 percent phenol solution was added to
each sample to react with the excess bromine. Samples were kept in closed
flasks and chilled in an ice bath to minimize loss of volatile forms of
cyanide. Four solvents, 3-pentanbiie, methyl isobutyl ketone, isopropyl
ether, and propionaldehyde, were evaluated for the extraction of cyanogen
bromide.
The percent recovery and standard deviation as determined by liquid
scintillation counting for extraction of three replicate samples by each
solvent are listed in Table 5. Recoveries ranged from 34 to 55 percent.
Aliquots of the sample solution taken after the addition of the reagents
were subjected to liquid scintillation counting to determine the amount of
cyanide lost' during the reaction. Only 4 percent of the cyanide was lost
from the sample. Thus, the low recoveries of cyanide are the result of low
extraction efficiency, not loss of cyanide from the sample.
Aqueous samples of unlabeled potassium cyanide were brominated and
extracted with methyl isobutyl ketone. These extracts were analyzed using
an electron capture gas chromatograph equipped with a column packed with
Porapak Q, 80-100 mesh. The oven temperature was maintained at 110°C.
Levels of cyanogen bromide as low as 1 ng on column were detected by this
-technique. Numerous problems, including water in the extracts and coa-
tamination of the methyl isobutyl ketone, were encountered.
22
-------
TABLE 4. RESULTS OF EVALUATION OF PROCEDURES FOR DETERMINATION OF CYANIDE
Method
Range
(mg/L)
Average +
standard
deviation
(cg/L)
Average
percent
recovery
Ultraviolet spectrophotometric quantification of the
tetracyanonickelate (II) anion complex formed by
reaction of hydrogen cyanide with ammoniacal nickel
chloride.
0.2 - 10
3.7 + 0.4
94
Spectrophotometric determination of the thiol anion 0.11-3.0 1.5+0.04 84
which is displaced by reaction of 5,51-dithiobis
(2-nitrcbenzoic acid) with cyanide ion.
Colorimetric determination based on Konig synthesis of 0.02 - 0.60 0.2 +.0.2 97
- a dye by the reaction of cyanide with chloramine-T to
form cyanogen chloride which subsequently reacts with
pyridine-barbituric acid reagent.
Colorimetric determination based upon formation and ex- 0.2 -1.0 1.0+0.1 109
traction of the neutral dicyano-bis(l,10-phenanthroline)
iron (II) complex produced through the exchange reaction
between tris (1,10-phenanthrolice) iron (II) and cyanide
ions.
Ultraviolet spectrophotometric determination based upon 0.2 -4.0 2.0+0.14 88
the absorbance of the chloranilate ion formed by reaction
of mercuric chloranilate with cyanide.
Determination of cyanide ion activity using an ion selective
electrode.
Direct 1 - 100 10+1 96
Indirect 0.06 - 1.00
v.
-------
TABLE 5. PERCENT RECOVERY OF CYANIDE
BY SOLVENT EXTRACTION AS CYANOGEN BROMIDE
Solvent
Percent recovered
+ standard
deviation
3-pentanone
methyl isobutyl ketone
isopropyl ether
propionaldehyde
50 + 2
55 + 0
34 + 1
47 + 2
The most sensitive quantification procedure is colorimetric determi-
nation based on the reaction of cyanogen chloride with pyridine-barbituric
acid reagent (Table 4). The pyridine-barbituric acid reagent procedure is
easy to conduct, and the only instrumentation required is a spectrophoto-
meter. The reaction of cyanide with cbloramine-T to produce cyanogen chlo-
ride and the subsequent reaction with pyridine-barbituric acid reagent to
form the dye are dependent on the temperature and pH of the sample solution.
Several compounds, particularly sulfide, interfere with the determination.
The remainder of the colorimetric determination procedures are not as sensi-
tive as the pyridine-barbituric acid procedure. The cyanide ion-selective
electrode provides a fast procedure for quantifying cyanide in aqueous
solutions, but it is subject to interferences and has a relatively high
detection limit.
Selection of Method
Distillation with stirring followed by quantification by the pyridine-
barbituric acid procedure was selected as the best method for determination
of cyanide in solid and semisolid samples. Stirring the sample with a
magnetic stirrer during distillation allows a larger size sample to be used
and reduces bumping of the distillation flask encountered with certain solid
matrices. Whether or not stirring is necessary depends on the nature of the
matrix and the size of sample. The expected cyanide concentration and the
desired detection limit should be considered when deciding on the appropri-
ate size of sample for distillation. The pyridine-barbituric acid colori-
metric procedure is sensitive to low concentrations of cyanide and is sub-
ject to minimal interferences.
PHASE III: OPTIMIZATION OF THE METHOD
During optimization studies, the most promising isolation and quantifi-
cation procedures for cyanide were joined to form a single method for analy-
sis of solid and semisolid environmental samples Jor cyanide. Cyanide was
isolated from the solid or semisolid sample matrix by distillation. The
cyanide in the distillate was quantified by the pyridine-barbituric acid
.24.
-------
procedure. The response of the quantification procedure to volatile sulfur
compounds, expected to be the most significant interference (82), was inves-
tigated. An absorber tube containing a lead acetate solution was utilized
for removal of volatile sulfur-containing compounds from the distillate.
Removal of the interferences and recovery of the cyanide were monitored
- using the pyridine-barbituric acid colorimetric procedure and the cyanide
ion-selectivc electrode. After experimental conditions had been optimized,
replicate samples of fish tissue were spiked with cyanide and analyzed to
determine the precision and accuracy of the method. This information and
the optimized method are presented below.
The method developed for analysis of solid and semisolid environmental
samples for cyanide consisted of distillation of hydrogen cyanide from an
acidified blurry of the sample into sodium hydroxide solution followed by
quantification of the cyanide in the solution by the pyridine-barbituric
acid colorimetric method. Sulfur-containing compounds which are distilled
from the sample and absorbed by the sodium hydroxide solution are known to
interfere with the colorimetric determination (82). The use of a trap
filled with lead acetate solution (83) was evaluated for removal of volatile
sulfur compounds from the distillate. A trap consisting of an open-ended
glass tube inserted into a tube containing 25 mL of 0.08M lead acetate
solution was placed in series between the distillation flask and the absor-
ber tube containing sodium hydroxide solution (Figure 2).
Samples of wet ground fish tissue weighing 10 g were placed in distil-
lation flasks, moistened with sodium hydroxide solution to insure basic pH
conditions, and spiked with 1 mL of a sodium hydroxide solution containing
100 Mg/L °£ potassium cyanide. The spiked samples were held for approxi-
mately 16 hours at A-6°C. Immediately prior to distillation, 500 mL of
distilled water, 50 mL of concentrated sulfuric acid, and 20 mL of magnesium
chloride solution were added to each sample. The samples were boiled for
2 hours to distill the cyanide which was absorbed in 35 mL of 1.8M sodium
hydroxide solution. Stirring the samples during distillation was not neces-
sary because of the reduction in sample size from 25 g to 10 g. The cyanide
in the distillate was quantified by both the pyridine-barbituric acid color-
imetric procedure and the cyanide ion-selective electrode. The colorimetric
procedure gives a negative response to sulfur compounds, and the. cyanide
ion-selective electrode gives a positive response. The two sets of results
were compared to detect the presence of interferences.
The percentages of cyanide recovered from samples of spiked fish dis-
tilled with and without the presence of lead acetate traps are presented in
Table 6. The average recoveries of cyanide from samples of fish tissue
distilled in the presence of traps containing lead acetate solution as
determined by the ion-selective electrode and the pyridine-barbituric acid
procedure are comparable. The average recovery of 54 percent is similar to
the recovery from fish tissue previously obtained. The average recovery of
cyanide from samples distilled without the use of lead acetate solution-
filled traps was determined to be 295 percent by the electrode and 25 per-
cent by the colorimetric procedure. This difference is probably due to the
f-presence of sulfur compounds in the distillate. Thus, it is necessary to
25
-------
I
Figure 2. Cyanide distillation apparatus equipped with
a lead acetate-filled absorber tube.
26
-------
use the lead acetate traps when distilling solid environmental samples to
remove potential interferences.
TABLE 6. COMPARISON OF PERCENTAGES OF CYANIDE RECOVERED FROM FISH TISSUE
SPIKED WITH 100 MICROGRAMS OF KCN DISTILLED WITH AND WITHOUT A LEAD
ACETATE TRAP AS DETERMINED BY A CYANIDE ION-SELECTIVE ELECTRODE AND
THE PYRIDINE-BARBITURIC ACID PROCEDURE
i Ion- Pyridine-
Replicate Lead acetate selective barbituric acid
number traps used electrode procedure
1 yes 51 50
2 yes 43 35
3 yes 78 89
4 yes 47 35
5 yes 56 55
average + standard deviation 55 + 14 53 + 22
6 no 77 26
7 no 295 43
8 no 514 5
average + standard deviation 295 + 218 25 + 19
Experiments were performed to determine the precision and accuracy of
the method for analysis of solid and semisolid environmental samples for
cyanide. Seven replicate 10-g portions of fish tissue were distilled, and
the cyanide concentration was determined by the optimized method. Three
groups consisting of seven 10-g replicates of the fish tissue were spiked
with 10, 25, and 100 |jg of cyanide as potassium cyanide. The samples were
analyzed, and the results are presented in Table 7. The average percent
recoveries were 94 ± 1.4, 95 ± 1-2, and 82 ± 5.6, respectively, for samples
spiked with 10, 25, and 100 pg of cyanide. The increase in recovery can be
attributed to improved technician skill.
The sensitivity of the method is dependent'-- on the size of sample dis-
tilled and is based on the concentration of cyanide which has an absorbance
reading of 0.02 at 578 nm. A detection limit of 0.5 pg/g can be achieved by
using a 10-g sample.
The method for analysis of solid and semisolid environmental samples
for cyanide is presented in Appendix B. Briefly, solid samples are blended
-or ground and placed in a distillation flask. Magnesium chloride solution
:.27.
-------
•and sulfuric acid are added to the sample, and hydrogen cyanide is distilled
into an absorber tube containing sodium hydroxide solution. An absorber
tube containing lead acetate solution is placed in series between the dis-
tillation flask and the absorber tube. The sample can be stirred while the
distillation is being performed using a combination magnetic stirrer-hot
plate. Stirring reduces bumping and, therefore, makes the use of a larger
sample feasible. The cyanide absorbed by the sodium hydroxide solution is
quantified by the pyridine-barbituric acid colorimetric procedure.
TABLE 7.! MICROGRAMS OF CYANIDE RECOVERED FROM SPIKED AND
UNSPIKED SAMPLES OF FISH TISSUE
Replicate
number
pg of cyanide
added to sample
0
10
25
100
1
0
i 9
23
75
2
0
7
23
80
-3
0
11
23
90
A
0
9
23
77
5
0
10
23
82
6
0
11
25
89
7
0
9
26
82
average pg recovered
0
9.4
23.7
82.1
standard deviation
0
1.4
1.2
5.6
coefficient of
15%
5%
7%
variation
percent of phenol
94%
95%
82%
recovered
PHASE IV: VALIDATION OF THE METHOD
The method developed for analysis of solid and semisolid environmental
samples for cyanide was validated by analysis of a variety of matrices. The
method consists of two steps: distillation of hydrogen cyanide from an
acidified slurry of the solid or semisolid sample and quantification of the
cyanide by the pyridine-barbituric acid procedure. Volatile
sulfur-containing compounds which might interfere with quantification of
cyanide are removed by bubbling the distillate through a solution of lead
acetate. Details of the method are presented in Appendix B.
The method for determination of cyanide in solid and semisolid environ-
mental samples was validated by the analysis of samples of ten matrices:
-28.
-------
1. bottom sediment from a lake
2. industrial sludge
3. industrial solid wastes from a disposal site
4. fish tissue
5. microinvertebrates (shrimp)
6. algae
_7_._ soil contaminated by chemicals from the disposal of industrial
sludges
8. growing vegetation
9. slags from industrial processing which are disposed of in a land-
fill i
10. fly ash from a commercial incinerator which is disposed of in a,'
landfill
One sample of each matrix was obtained, and six 10-g portions of each
sample were frozen to preserve the cyanide. Two portions of each sample
matrix were analyzed by the developed method to determine the concentrations
of cyanide in the samples. The four remaining portions of each matrix were
removed from the freezer, spiked with a sodium hydroxide solution containing
potassium cyanide, and analyzed. Two portions of each matrix were spiked
with 26 pg of cyanide, and the remaining two were spiked with 80 or 200 pg
of cyanide to give spiked cyanide concentrations of 2.6, 8.0, and 20 pg/g,
respectively. Some of the environmental samples contained cyanide, and,
thus, the total cyanide concentration of these samples was greater than the
spiked concentration alone (Table 8).
The concentrations of cyanide recovered from each sample are presented
in Table 8. The recovery of cyanide from the AO spiked samples ranged from
69 to 112 percent, with an average of 90 ± 10 percent. Recovery did not'
vary with the concentration of cyanide over the range of 2.6 to 20 pg/g.
Twenty samples were spiked with 2.6 pg/g of cyanide. The. average percent
recovery of cyanide from these samples was 90 ± 10. Average percentages of
cyanide recovered from 12 samples spiked with 8.0 pg/g and eight samples
spiked with 20 pg/g were 89 ± 13 and 90 ± 11, respectively.
The quality of the data was verified by a rigid quality control pro-
gram. A calibration curve was prepared each day using standard solutions of
potassium cyanide. These standards were not distilled. A reagent-glassware
blank consisting of 500 mL of distilled water was distilled and analyzed
daily. Performance of the distillation apparatus and quantification proced-
ure was checked daily by distillation and analysis of one quality control
sample for each group of four environmental samples. The quality control
samples consisted of 500 mL of distilled water spiked with a solution of
potassium cyanide to give a concentration of 50 to 400 pg/L. The recoveries
of cyanide from the quality control samples are presented in Table 9. The
average percent of cyanide recovered from 500-mL volumes of distilled water
containing from 25 to 200 pg of cyanide was 88 ± 7. This compares well with
the average percent recovery of 89 ± 10 from the 40 spiked environmental
samples. ; i
29,
-------
TABLE 8. CONCENTRATIONS OF CYANIDE ADDED TO AND RECOVERED
FROM SOLID AND SEMISOLID ENVIRONMENTAL SAMPLES
Average
— Concentration
Concentration
Percent
percent
Matrix added (pg/g)
recovered (|Jg/g)
recovery
- recovery
bottom sediments 0
ND
84 + 6
0
ND
2.6
2.3
88
2.6
2.0
77
8.0
7.2
90
8.0
6.4
80
industrial sludge 0
1.8
84 + 12
0
1.4
2.6
3.6
77
2.6
4.2
.100
20
16
72
20
19
87
industrial solid 0
ND
94 + 11
waste 0
ND
2.6
2.3
88
2.6
2.3
88
20
18
90
20
22
110
industrial 0
ND
94+3
processing slag 0
ND
2.6
2.5
96
2.6
2.5
96
20
19
95
20
18
90
incinerator fly ash 0
0.9
95+7
0
1.0
2.6
3.6
100
2.6
3.6
100
20
18
85
20
20
95
contaminated soil 0
ND
103 + 12
0
ND
2.6
2.2
85
2.6
2.9
112
8.0
8.6
108
8.0
8.6
108
30
-------
Table 8 (Continued).
Matrix
t
Concentration
added (pg/g)
Concentration
recovered (pg/g)
Percent
recovery
Average
percent
recovery
fish tissue
. 0
ND
0
ND
2.6
2.5
96
2.6
2.3
88
8.0
8.2
102
8.0
7.3
91
microinvertebrates
0
ND
(shrimp)
0
ND
2.6
2.3
88
2.6
2.6
100
8.0
7.4
92
8.0 .
6.9
86
algae
0
ND
0
ND
2.6
2.0
77
2.6
2.0
77
8.0
6.1
76
8.0
6.1
76
vegetation
0
ND
0
ND
2.6
2.3
88
2.6
2.3
88
8.0
7.1
89
8.0
5.5 .
69
94 +
76 + 1
84 + 10
ND = not detected above detection limit of 0.5 pg/g
31
-------
TABLE 9. RECOVERY OF CYANIDE FROM AQUEOUS QUALITY ASSURANCE SAMPLES
Percent
Sample pg of cyanide in 500 ml (Jg of cyanide recovered recovery
.25
-20 -
80
2
26
25
96
3
26
22
85
4
26
26
100
5
200
174
87
6
200
158
79
7
26
23
88
8
80
75
94
9
26
25
_ 96
10
80
68
85
11
80
72
90
12
26
22
85
13
26
24
92
14
80
60
75
15
80
67
84
average + standard deviation 88+7
32.
-------
SECTION 4
METHOD FOR ANALYSIS OF SOLID AND SEMIROLTD ENVIRONMENTAL SAMPLES FOR TOTAL
PHENOLIC COMPOUNDS
INTRODUCTION
The literature was searched for methods for analysis of any matrix for
total phenolic compounds. Numerous publications were located using com-
puterized and manual literature searching techniques. The majority of the
references discussed analysis of aqueous samples. Limited information was
found on analysis of solid or semisolid matrices. During Phase I, methods
which were appropriate for application to the analysis of solid and semi-
solid environmental samples were selected for empirical evaluation. Selec-
tion criteria used in evaluating the methods in the literature were accuracy
and analytical range of the methods, lack of significant interferences,
minimal sensitivity to changes in sample pH or temperature, and ease of
performing the determination.
Evaluation of methods of analysis for total phenolic compounds was
approached as a two-step procedure. First, the phenolic compounds were
isolated from the solid or semisolid matrix. Solvent extraction and steam
distillation were the isolation procedures evaluated. Radiolabeled phenolic
compounds were added to solid and semisolid samples, and recovery by the
isolation procedures was monitored using liquid scintillation counting.
Once the phenolic compounds are in a solution, the concentration can be
determined. Three quantification procedures were tested:
1. Phenol + 4-aminoantipyrine •+ antipyrine dye
2. Phenol + 3-methyl-2-benzothiazolinone hydra zone -»¦ azo dye
3. Bathochromatic shift occurring upon ionization of the OH-grovip
of phenol
The quantification procedures were evaluated by analysis of solutions oi
phenol in distilled water. The sensitivity, analytical range, precision,
and accuracy of each method were compared. Solvent extraction was selected
as the isolation procedure because it recovers a greater percentage of
phenolic compounds than distillation. Measurement of the change in absor-
bance due to ionization of the hydroxyl group of phenol using the ultra-
violet ratio spectrophotometer most sensitive procedure for phenol in dis-
! tilled water. However the 4-aminoantipyrine method was chosen because it
has been shown to work on a large variety of aqueous environmental matricesr
and the results can be compared to an abundance of accumulated data.
In the optimization phase, solvent extraction and the 4-aminoantipyrine
procedure were joined to form a method for analysis of solid and semisolid
33
-------
environmental samples for total phenolic compounds. Experiments were per-;
formed using environmental samples spiked with phenol to determine optimal
analytical conditions.
The method was validated by analysis of samples of ten solid or semi^
solid environmental matrices. The samples were spiked with two levels of
phenol and analyzed. Precision, accuracy, and sensitivity of the method are
discussed below.
PHASE I: SELECTION OF METHODOLOGY FROM THE LITERATURE
The literature was searched for methods of analysis of any type of
matrix for phenolic compounds. Computerized retrieval techniques were used
for searching two data bases, CA CONDENSATES/CASIA and TOXLINE. Chemical
abstracts for the years 1967 to 1970 and Government Reports and Announce-
ments Index published by the National Technical Information Service for the
years 1968 to 1977 were searched manually. Approximately 71 papers of
interest were identified by these methods. References cited in the reviewed
literature were also obtained.
Methods of Isolation
Samples which are to be analyzed for phenolic compounds frequently
contain other materials which may contribute to the turbidity or color of
the sample or interfere with the determination in other ways. Three basic
techniques are used to separate phenolic compounds from an aqueous matrix.
These are distillation, solvent extraction, and liquid chromatography.
Steam Distillation—
Distillation of wastewater samples prior to analysis is recommended by
the U.S. EPA (9) and the American Public Health Association (13) to separate
the phenolic compounds from nonvolatile components of the sample which could
interfere with the determination. The phenolic compounds are distilled
from an acidified sample at a constant rate. Phenolic compounds volatilize
gradually; therefore, the entire sample must be distilled to ensure complete
recovery of all the phenolic compounds (9). Recovery of 80 and 90 percent
of the volatile phenolic compounds during steam distillation and normal
distillation, respectively, has been reported (84). While distillation
serves to separate phenolic compounds from nonvolatile components of the
sample, it does not concentrate the phenolic compounds.
Solvent Extraction—
Solvent extraction can be used to separate and concentrate phenolic
compounds from the sample matrix. Solvents which have been used to extract
phenols from aqueous solutions include dichloromethane (85), petroleum ether
(86), chloroform (87,88), toluene (89), dimethylformamide (90), and tributyl-
phosphate (91). A potassium hydroxide solution has been used to extract
phenolic compounds from petroleum products (92). To concentrate the phe-
nolic compounds further, the pH of the caustic extract is adjusted to less
34
-------
than seven and the phenols are extracted with dimethylether. The dimethyl-
ether is evaporated, and the residue is dissolved in acetic acid. Refinery
wastewater samples at pH 12 have been extracted with carbon tetrachloride to
remove oil and then extracted at pH 5 with tributylphosphate to recover the
phenols as a concentrate (91). Phenolic compounds can be separated from
other organic compounds in wastewater by converting the phenols to their
salts by the addition of sodium hydroxide (93). The organic compounds are
removed from the sample by extraction with a 1:1 mixture of ether-petroleum
ether. After separation of the phases, the aqueous phase is acidified by
addition of a mineral acid to convert the phenolic salts to free phenols.
Liquid Chromatography—
Ion-exchange, adsorption, and gel chromatography can be used to sepa-
rate phenolic compounds from a sample matrix. The macroreticular anion
exchange resin Amberlyst A-26 and the porous polystyrene-divinyl benzene
copolymer XAD-2 (94), as well as the cation exchange resins Amberlite
IR 112, Amberlite IRC 50, and Dualite C 25 (95), have been used to adsorb
phenolic and other organic compounds from aqueous samples. A method for
separating phenolic compounds from a sample matrix based on adsorption of
phenolate ions from alkaline solution on the A-26 macroporous anion-exchange
resin in the hydroxyl form was developed (96). The adsorbed phenolate ions
are converted to the molecular form by washing the column with hydrochloric
acid and are eluted from the column with an acetone-water solution. The
average recovery of eight phenols from a spiked tap water sample was
97.7 percent (96).
Phenolic compounds can be removed from water by adsorption to carbon or
Sephadex gels (97). Because the adsorption capacity of activated carbon is
limited, this technique is better suited to sampling water which is clean
than water which contains high concentrations of organic compounds (98).
The usefulness of the carbon adsorption technique is limited by the adsorp-
tion efficiency of 94 percent and desorption efficiency of 22 percent (99).
Sephadex gels effectively separate free phenols from high molecular weight
proteins and phenolic polymers (100).
Methods of Quantification
Spectrophotometric Methods—
A method for quantification of total phenolic compounds should be able
to determine ortho-, meta-, and para-substituted phenolic compounds. The
primary shortcomings of many colorimetric methods are that they lack sensi-
tivity to some phenolic compounds (101,102) and produce complexes which
absorb at different wavelengths (92,103-105). The five most widely used
methods for determining trace levels of phenolic compounds in water were
reviewed (106), and it was concluded that the^-aminoantipyrine procedure is
the fastest, most precise, and most accurate when compared to the following
spectrophotometric methods: Gibbs1 dibromoquinone chlorimide, nitroso-
phenol, infrared, and ultraviolet.
The 4-aminoantipyrine method is currently the most widely recommended
method for analysis of water and wastewater samples (9,13). It is based on
-------
the formation of antipyrine dyes by the reaction of phenolic compounds with
4-aminoantipyrine in the presence of an alkaline oxidizing agent (107-109).
Compounds which have an open para position or which have groups such as
halogen, carboxyl, sulfonic acid, hydroxyl, or methoxyl in the para position
are detected (106). Phenolic compounds in which the para position is
blocked by an aryl, alkyl, nitro, nitroso, benzoyl, or carbonyl group are
'not determined by this method. The dyes produced by all phenolic compounds
have a maximum absorbance at the same wavelength. The method has been ap-
plied to determination of total phenolic compounds in wastewater (106,110)
and surface water (111), 2,4-dichlorophenol in water (86), 1-naphthol in
aqueous solutions of carbaryl (88), and phenolic fungicides in fabric (112).
The classical method of analysis for phenolic compounds is based on the
formation of dibromoindophenol dyes by the condensation of 2,6-dibromoqui-
none chlorimide with phenolic compounds having an open para position in
alkaline solution (113). The relationship between concentration of a phe-
nolic compound and absorbance of the dye is linear over the range 0 to 10
mg/L in aqueous solution or 0 to 100 |Jg/L in n-butyl alcohol. The time
required for dye formation is dependent on the phenolic compounds being
determined with up to 24 hours required for complete color development of
some compounds. Another inherent problem is that the reaction products of
different phenolic compounds in combination with this colorimetric reagent
have absorption maxima at different wavelengths (106,114). An advantage of
this is that specific phenolic compounds such as resorcinol can be determin-
ed in the presence of phenol by meaGuring the absorbance of the dibromoindo-
phenol dye at 530 nm immediately after the addition of reagents to the
sample (115).
A method for determination of phenolic compounds based on the for-
mation of indophenol by the reaction with N-(benzenesulfonyl) quinonimine
was reported (116). The reagents used are much more stable than 2,6-dibro-
moquinone chlorimide, and color formation is complete within three minutes.
The concentration of phenolic compounds is proportional to the initial rate
of formation of indophenol and to the absorbance of indophenol after three
minutes. Unfortunately, the wavelength of maximum absorbance ranges from
610 to 650 nm depending on the specific phenolic compound. Lowest detect-
able concentrations range from 0.6 to 10.0 mg/L for seven phenols deter-
mined .
A promising method for quantification of total phenolic compounds which
has been automated (117,118) is based on the oxidative coupling reaction of
3-methyl-2-benzothiazolinone hydrazone with phenolic compounds in an acid
solution of eerie ammonium sulfate to from an azo dye. The reaction is
relatively insensitive to pH, produces a more intensely colored product than
the 4-aminoantipyrine procedure, and occurs with ortho-, meta-, and para-
substituted phenolic compounds. Aromatic amines and aliphatic aldehydes
react to give a falsely high result if not removed from the sample prior to
analysis (117).
Most spectrophotometric absorption methods depend on the reaction of
the aromatic ring of phenol with reagents to produce a compound with a high
-------
-molar absorptivity. Ionization of the phenol hydroxyl group when the pH is'
shifted from acidic to basic is the basis of spectral UV methods. Sensitive
methods for quantification of phenolic compounds based on this bathochro-
matic shift of the long wavelength maximum which occurs when phenols are
made basic have been developed for analysis of refinery wastes (91,119),
synthetic rubber latex and rubber products (120), and serum (121). A method
:was proposed for quantifying phenolic compounds by monitoring the absor-
bance at a wavelength of 292.5 nm (119). This wavelength was selected
because it is an intermediate value for the range of absorption maxima
exhibited by a group of representative phenolic compounds. P-cresol was
selected as the calibration standard because it has an absorption maximum at
292.5 nm which is similar to the absorption maximum of a selected group of
phenols. The shift of the wavelength maximum can be quantified by'measuring
the absorbance before and after pH adjustment with a standard single or
double beam ultraviolet spectrophotmeter or with an ultraviolet ratio spec-
trophotometer specially designed for this technique (122,123). Phenol,
rather than £-cresol, is used as the standard with the ultraviolet ratio
spectrophotometer. The method is sensitive to all except hindered phenolic
compounds which do not ionize fully,.such as those having a butyl or larger
group in the 2 or 6 position (91). Concentrations of phenolic compounds as
low as 0.25 mg/L were determined by differential absorbance measurements
using a double beam ultraviolet spectrophotometer with 5 cm absorption cell
(118).
Titrimetric Methods—
There are many suitable titrants and solvents (124) for volumetric
determination of phenolic compounds. Titrants- uied in most cases are not
specific for phenolic compounds. Examples of these are the oxidimetric
titrants N,N' -dibromo-p-toluenesulphonamide (dibromamine-T) (125) and iodo-
benzene dichloride (126) which are used in acetic acid medium for potentio-
metric quantification of many diverse reductants and the titrant 1,2,4,6-
tetraphenylpyridinium acetate which is used for potentiometric precipitation
titration of nitrophenols, dinitrophenols, and some halogenated phenols, as
well as nitroform and nitroform yielding compounds (127). Many volumetric
determinations are based on the bromination of phenolic compounds (126,128-
131). These methods are suitable for quantification of a specific phenolic
compound in a solvent containing no other compounds that react with the
titrant. Catalytic thermometric titrations (132-135) can be used for quan-
tification of phenolic compounds, but the methods generally determine all
weak acids, thus requiring the separation of phenols from the sample matrix
prior to titration. Titration methods were not considered for evaluation
because they lack sensitivity or require special instrumentation.
Chromatographic Methods—
Chromatography has been used extensively to separate and identify
phenolic compounds. Gas chromatography (136-139) and capillary column
chromatography (140) with electron capture (141) or flame ionization detec-
tors (90,123,137,142) are used to quantify phenolic compounds. Researchers
have prepared chloroacetate (141), O-isobutyloxycarbonyl (142), chloro-
phenyl-2,4-dinitrophenyl ether (139), and diethyl phosphate (143) derivia-
tives of phenolic compounds to improve sensitivity. Unlike many of the
-------
"spectrophotometric methods, chromatographic methods are sensititive to all
phenolic compounds, but the specificity achieved by chromatography is not
required for determination of total phenols. Chromatographic methods are
applicable to determination of individual phenolic compounds rather than
total phenolic compounds; therefore, they were not considered for evalu-
ation. — ¦
Selection of Methods
Methods which appeared to be applicable to the analysis of solid and
semisolid environmental samples for total phenolic compounds were selected.
The separation technique of choice depends on the matrix being analyzed, the
phenolic compounds present, and the expected concentration of the phenolic
compounds. The separation technique must be compatible with the determi-
nation step which follows. Distillation is a suitable separation method for
aqueous samples containing few volatile compounds which could interfere in
the determination. The distillation procedure is time consuming but easy to
conduct. Extraction and adsorption techniques are useful for concentrating
phenolic compounds as well as separating them from the sample matrix. Tha
recovery of a variety of phenolic compounds must be determined to verify the
usefulness of the separation technique as part of a method for determination
of total phenolic compounds. Solvent extraction and steam distillation were
the isolation techniques selected for evaluation.
Selection of procedures for the quantification of phenolic compounds
was based on analytical response to the phenolic compounds, accuracy and
precision over a wide concentration range, stability of reagents used for
the determination, and ease of manipulation. The following methods for
determination of total phenolic compounds were evaluated.
1. Spectrophotometric determination based on measurement of the
change in absorbance exhibited' when the wavelength of maximum
absorbance shifts due to a change in pH of the sample solution
(119) using the ultraviolet ratio spectrophotometer (122, 123).
2. Coloriraetric determination by measurement of the absorbance of the
dye produced by the oxidative coujling reaction of 3-methyl-2-
benzothiazolinone hydrazone with phenolic compounds (117).
3. Colorimef.ric determination by measurement of the absorbance of the
dye produced by the reaction of 4-aminoantipyrine with phenolic
compounds (9).
\
PHASE II: EVALUATION OF THE METHODS
\
The purpose of this phase was to eliminate nonviable methods and iden-
tify necessary modifications of promising procedures; Development and evalu'
ation of methods was approached as a two-step procedure. Separation and
determination of phenolic compounds were treated independently. Two isola-
tion techniques, steam distillation and solvent extraction, were evaluated
by application to solid and semisolid environmental samples spiked with
1 .¦
38
-------
phenolic compounds. Three determination procedures were tested on samples
of distilled water containing phenol. j
Three types of solid or semisolid environmental matrices were selected'
to be test samples from the list of ten of prime interest. The samples were
selected to cover the range of characteristics expected in environmental
matrices. A pool of fish tissue was prepared by chopping the edible por-
tions of fish into small pieces, homogenizing the tissue in a olender, and
freezing it until needed. Lake sediment, consisting of fine and medium
sized sand grains, was dried at 103°C and then separated from miscellaneous
material such as shells, leaves, and sticks. Domestic sludge was obtained
from the drying pond of a sewage treatment plant. The sludge, dried at
103°C, was pulverized with a mortar and pestle. The dried, pulverized
sludge was stored at room temperature until used.
Three phenolic compounds—phenol, £-nitrophenol, and 2,4,5-trichloro-'
phenol—were selected for use as model compounds. Selection was based on
probability of occurrence in environmental ^amples, chemical characteristics
of the compound, and availability of the C-labeled isotopes. The use of
radiolabeled compounds permitted evaluation of the efficiency of the sepa-
ration procedures independently of procedures specific for the determination
of phenolic compounds.
i
t
Procedures for separation and determination of phenolic compounds were
evaluated by performing the procedure on seven replicate samples. Percent
recovery, average percent recovery, and standard deviation of the replicates
were calculated using the following equations: I
| i
i
! ¦ i
100 (0.) I
Percent recovery = ^ ]
|
where 0^ = observed value j
T, = true value !
1 !
i n
i P.
Average percent recovery = i=l
n
where n = number of samples
|
H
N-l
= percent recovery of sample i
Standard deviation =
_ Ap< • (I *
-------
Procedures for Isolation
Two types' of isolation procedures, solvent extraction and steam dis-
tillation, were applied to the separation of phenolic compounds from solid
and semisolid environmental samples. These procedures were evaluated by
-application to samples spiked with radiolabeled phenolic compounds. Samples
'of -a solid matrix were spiked with known concentrations of radiolabeled
phenol, j>-nitrophenol, or 2,4,5-trichloroplienol. Spiking was tccomplished
by adding a small volume of a solution containing the phenolic compound to
the matrix which had been moistened with frater to facilitate mixing. For
the first several experiments, samples were held for 1 hour after spiking
before being treated by a separation procedure. In the majority of experi-
ments, however, samples were allowed to sit for 16 hours to permit adsorp-
tion of the phenolic compounds to the matrix.
Controls were prepared with each set of solid samples and consisted of
distilled water or other solvents spiked with the same concentrations of
radiolabeled model phenolic compounds. The spiking solution and the control
samples were subjected to liquid scintillation counting by transferring
three replicate 0.5 or 1.0 mL aliquots of the solution to a glass vial
containing 10 mL of scintillation cocktail. The counts obtained for the
samples from a liquid scintillation counter were corrected for counting
efficiency and used to calculate the concentration of phenolic compounds in
the sample solutions.
Solvent Extraction—
Two techniques of solvent extraction were investigated. The first
technique consisted of stirring the matrix with a solvent using a magnetic
stirring bar. Experiments were performed using water at pH 2 as the solvent
and also using methylene chloride as the solvent. A 10-g portion of dried
sludge was placed in a 250-mL glass bottle, spiked with 2 pg radiolabeled
phenol, and allowed to air dry overnight. One hundred milliliters of a
solvent was added to the sample in the bottle, and the mixture was stirred
with a magnetic stirring bar. Periodically, 5-mL aliquots of the solvent
were removed from the bottle and subjected to liquid scintillation counting
to determine the recovery of the phenolic compound. The ratio of solvent to
sludge decreased with time because of the removal of aliquots for counting.
Details of the experiment and percent recovery of phenol are shown in
Table 10. Recovery using water acidified with sulfuric acid to pH 2 for the
extraction solvent was low as expected. Extraction with methylene chloride
yielded a higher recovery. Recovery remained fairly constant over time.
i
Another set of experiments was conducted using the same procedure, but
the volume of solvent removed for liquid scintillation counting, usually
5 mL, was replaced each time with fresh solvent. Recoveries of the phenolic
compounds were calculated from liquid scintillation counting data and took
into account the dilution effect of adding fresh methylene chloride at each
time interval. The average recoveries of phenol, j>-nitrophenol, and 2,4,5-
trichlorophenol extracted from seven replicate samples of fish tissue,
sediment, and sludge are listed in Table 11. Recovery tended to be highest
after 2 hours of stirring. The average percentage of phenol recovered from
-------
TABLE 10. AVERAGE PERCENT PHENOL EXTRACTED FROM SLUDGE BY THE
STIR-BAR EXTRACTION PROCEDURE WITHOUT VOLUME REPLACEMENT
Solvent
pH of
mixture
Extraction
time (hour)
Average percent
recovery ±
standard deviation
Number
of
replicates
water
.2
0
38
+
6
7 \
i
0.5
41
+
5
l
n 1
1.0
28
+
5
7
i
2.0
24
±
8
7 1
4.0
20
±
4
7 !
1
methylene
NA*
0
64
±
9
7
j
chloride
0.5
79
+
3
7
1.0
80
±
3
2.0
78
+
3
4.0
78
+
3
7 !
| * Not adjusted j
i i
j ' ; j
| all three matrices was 87. An average of 85 percent of the £-nitrophenol
j and 80 percent of the 2,4,5-trichlorophenol were extracted from the three
¦ matrices.
! . i
The second solvent extraction technique evaluated was Soxhlet extrac-
i tion with methylene chloride as the solvent. A series of experiments was
performed to determine the recovery of phenol, £-nitrophenol, and 2,4,5-tri-
chlorophenol from sludge, sediment, and fish tissue. Ten-gram portions of
; each solid matrix were placed in a cellulose extraction thimble and spiked
with a solution containing a known concentration (Table 12) of radiolabeled
, phenol, g-nitrophenol, or 2,4,5-trichlorophenol. The spiked samples were
i allowed to air dry overnight at room temperature. The samples were ex-
' tracted for two hours with methylene chloride in a Soxhlet extraction ap-
; paratus. Controls were prepared by extracting an empty cellulose thimble
| with methylene chloride containing one of the phenolic compounds to deter-
, mine losses of phenol to the extraction apparatus. Method blanks were
j prepared by extracting a cellulose thimble with-methylene chloride. Seven
' replicate extractions of each matrix spiked with each phenolic compound were
! performed. Aliquots of methylene chloride were analyzed by liquid scintil-
i lation counting to determine the amount of radiolabeled phenolic compound
¦' extracted. The average recovery and standard deviation of each combination
, of matrix and phenolic compound are shown in Table 12. The recoveries
[ranged from 52 to 99 percent and varied with sample matrix and phenolic
jv. compound. _ __
J 5 • i ....
41.
-------
TABLE 11. AVERAGE PERCENT RECOVERY OF PHENOLIC COMPOUNDS EXTRACTED FROM SEVEN REPLICATES OF TEN
GRAMS OF EACH MATRIX BY STIRRING WITH METHYLENE CHLORIDE FOR TIMES UP TO FOUR HOURS
Average percent recovered
+ standard deviation
pg of compound
Extraction time
(hr)
. .
added to sample
0
0.
.5
1.
.0
2.0
4.
0
phenol
tissue
1.3
16 + 3
72
+
6
86
+
9
93 + 9
94
+ 8
sediment
1.4
66 + 24
83
+
16
84
+
16
84 + 16
85
+ 13
sludge
1.5
65 + 5
86
+
3
86
+
3
86 + 3
88
+ 4
p-nitrophenol
tissue
3.4
14 + 2
65
+
4 :
69
+
6
73 + 6
72
+ 4
sediment
3.5
48 + 8
95
+
1 '
96
+
2
98 + 3
97
+ 3
sludge
2.0
30 + 7
72
+
* .
79
+
6
83 + 7
88
+ 8
2,4,5-trichloro
-
. . .
—
¦
... .
phenol
tissue
56.5
11 + 2
62
+
4 '
69
+
6
72 + 6
75
i + 4
sediment
31.0
32 + 5
90
+
8 i
94
+
2
97 + 2
95
i + 2
sludge
18.0
36 + 5
68
+
3 ,
71
+
4
72 + 3
75
1 t 5
-------
Matrix
/.t
14,
I
i <
t
TABLE 12. RECOVERY OF C-LABELED PHENOLIC COMPOUNDS FROM SLUDGE, SEDIMENT, AND
FISH BY SOXHLET EXTRACTION WITH METHYLENE CHLORIDE !
14
C-Phenol
14
14
C-p-nitrophenol C-2,4,5-trichlorophenol
Average
percent
Concentration recovery ±
in sample standard
(Mg/g) deviation
Concentration
in sample
(Mg/g)
Average
percent
recovery ±
standard
deviation
: Average
- • JjS* •
J percent
Concentration i recovery ±
in sample ' standard
ClJg/g) j deviation
Sludge
0.14
75 + 7
0.34
73 + 13
4.6
77 + 17
Sediment
0.27
90 + 1
0.67
99 + 9
2.5
: 91+10
t
Fish
0.28 56 + 5
0.11
52 + 6,
1.6 i -90 + 7
-------
Steam Distillation—
Steam distillation is the conventional procedure for separating phe-
nolic compounds from aqueous samples and was evaluated as a procedure for
separating phenolic compounds from solid and semisolid environmental sam-
ples. A series of experiments was performed to assess the recovery of
phenol, £-nitrophenol, and 2,4,5-trichlorophenol from samples of water and
'.sludge. . Ten-gram portions of dried, pulverized domestic organic sludge were
prepared for distillation by spiking with a solution of radiolabeled phenol,
p-nitrophenol, or 2,4,5-trichlorophenol. The samples were allowed to air
dry overnight at room temperature. One hundred milliliters of water was'
added to the samples, and the pH of the mixture was adjusted to 4 with'
phosphoric acid. The mixture was gently boiled to distill the water and
phenolic compounds. Aliquots of the distillate were counted by liquid
scintillation counting periodically to determine the recovery of the pheno-
lic compound. Control samples were prepared by spiking 100 mL of distilled
water with a phenolic compound and adjusting the pH to 4 with phosphoric
acid. Controls and sludge samples were prepared at the same time and were
treated identically. The average percent recoveries of phenol, £-nitro-
phenol, and 2,4,5-trichlorophenol distilled from controls and sludge samples
are shown in Table 13. Recovery of phenol from both the water controls and
the sludge samples was 92 to 96 percent. Recovery of j>-nitrophenol and
2,4,5-trichlorophenol was less than 5 percent.
TABLE 13. AVERAGE PERCENT RECOVERY OF PHENOL, P-NITROPHENOL,
AND 2,4,5-TRICHLOROPHENOL BY DISTILLATION OF AQUEOUS AND SLUDGE SAMPLES
Average percent
pg of recovered + standard Number of
Matrix and compound compound deviation rep]icates.
7
2
2
7
3
5
Procedures for Quantification
Three procedures for the quantification of total phenolic compounds,
included in Appendix C, were evaluated by analysis of aqueous solutions of
water
phenol
g-nitrophenol
2,4,5-trichlorophenol
Sludge
phenol
£-nitrophenol
2,4,5-trichlorophenol
402 92+5
402 <1
417 2+1
2 96 + 7
3 <1
31 <1
.44
-------
phenol. Seven replicate analyses were performed by each procedure on a
solution having a concentration approximately ten times that of the lowest
calibration standard. One of the procedures evaluated is based on the
reaction of phenolic compounds with 4-aminoantipyrine in the presence of
potassium ferricyanide at a pH of 10 to form a dye (9). The absorbance of
the solution containing the dye is measured at a wavelength of 460 or 510 nm
and is proportional to the concentration of phenolic compounds in the sam-
ple. The 4-aminoantipyrine reacts with the carbon ia the para-position of
the phenolic compound. Th'. reaction does not take place if the para-posi-
tion is occupied by a sut-stituent other than a carboxyl, halogen, hydroxyl,
methoxyl, or sulfonic acid group (144). Materials present in the sample
which cause turbidity or color interfere with the determination. The second
colorimetric procedure evaluated was based on measurement of the absorbance
of a dye produced by the oxidative coupling reaction of 3-methyl-2-benzothi-
azolinone hydrazone with phenolic compounds. The coupling takes place at
the ortho-position or at the para-position if the ortho-position is unavail-
able (145).
The analytical range, precision, and accuracy of these two procedures,
reported in Table 14, are very similar. The reagent 3-methyl-2-benzothia-
zolinone is able to react with a greater number of substituted phenolic
compounds than is 4-aminoantipyrine, and the phenol reaction product has a
higher molar absorptivity than the comparable 4-aminoantipyrine product. A
disadvantage of the 3-methyl-2-benzothiazolinone procedure is that the
compounds produced by reaction with different phenolic compounds have their
absorbance maxima at different wavelengths ranging from 460 to 595 nm (145).
The third procedure evaluated for the determination of total phenolic
compounds is based on the bathochromatic shift which occurs when the pH of
the sample solution is shifted from acidic to basic (119,122,123). The
shift in the wavelength of maximum absorbance permits measurement of the
phenolic compound concentration using a standard ultraviolet spectropho-
tometer or an ultraviolet ratio spectrophotometer developed specifically for
the purpose by Spt.ctro Products, Inc., New Haven, Connecticut. The detec-
tion limit achievable with the Spectro Products instrument is much lower
than that obtained using a standard ultraviolet spectrophotometer
(Table 14). P-cresol was selected as the calibration standard for use with
the standard ultraviolet spectrophotometer because it has a wavelength
maximum at 292.5 nm, which is similar to the mean wavelength maximum of a
selected group of phenolic compounds (ll'J). Phenol was selected for use as
the calibration standard for the Spectro Products instrument as recommended
by the manufacturer (146).
Selection of Method
The goals of the second phase were to evaluate procedures for isolation
and quantification of total phenolic compounds and select the best proce-
dures for further modification and development. The isolation procedure
yielding the highest recovery of phenol, g-nitrophenol, and 2,4,5-trichloro-
phenol from samples of fish tissue, sediment, and sludge was solvent extrac-
tion performed by stirring the sample with methylene chloride. The recov-
eries achieved by this procedure ranged from 72 to 98 percent.
-------
TABLE 14. RANGE, PRECISION, AND ACCURACY OF PROCEDURES FOR THE DETERMINATON
OF TOTAL PHENOLIC COMPOUNDS
Method
Range
(mg/L)
Average +
standard
deviation
(mg/L)
Average
percent
recovery
Colorimetric determination by measurement of the
absorbance of the dye produced by the oxidative coupling
reaction of 3-methyl-2-benzothiazolinone hydrazone (MBTH)
with phenolic compounds.
Colorimetric determination by measurement of the
absorbance of the dye produced by the reaction of
4-aminoantipyrine with phenolic compounds.
0.004 - 0.060 0.040 + 0.004
0.004 - 0.04 0.040 + 0.002
100
108
Spectrophotometric determination based on
measurement of the change in absorbance
when the wavelength of maximum absorbance
shifts due to a change in pH of the sample
solution. _
Sppctro Products PH-3 instrument
using phenol standards
sample containing phenol
Standard ultraviolet spectrophotometer
using jj-cresol standards
sample containing £-cresol
sample containing phenol
0.002 - 0.10
1.0 - 30
1.0 - 30
0.02 + 0.002
8.5 + 0.7
25 + 1
98
85
236
-------
Phenolic compounds can be extracted from the methylene chloride into'a
dilute solution of sodium hydroxide and then quantified by one of several
procedures. The most sensitive procedure for determination of phenol in
distilled water is the one based on measurement of the bathochromic shift of
phenolic compounds which occurs when the pH of the solution is changed from
4 to 12 using the ultraviolet ratio spectrophotometer. The procedure is
'dependent on ionization of the phenol hydroxyl group and independent of the
presence and position of substituent groups. Disadvantages of this proce-
dure are that phenolic compounds which exist as tautomers and certain other
phenolic compounds such as the highly substituted chlorinated phenolic
compounds are not detected because their absorption maxima lie outside of
the range for phenol and the phenolate ion. Absorbance can be determined
with an ultraviolet spectrophotometer or an ultraviolet ratio spectrophoto-
meter designed specifically for the determination of phenolic compounds.
The ultraviolet ratio spectrophotometer is approximately 500 times more
. sensitive than the standard ultraviolet spectrophotometer. Quantification
using an ultraviolet ratio spectrophotometer such as the one man>
-------
can be removed from storage, placed directly in the blender to be homog-
enized and then extracted. Sodium sulfate is added as a drying agent to the
frozen sample in the blender, and the mixture is blended to produce a dry
powdered sample. The sample is mixed with 100 mL of 0.18M sulfuric acid
solution, transferred to a centrifuge tube, and centrifuged to separate the
solids from the acid solution. The supernatant acid solution is decanted
into a separatory funnel and extracted three times with methylene chloride
using volumes of 50, 25, and 25 mL. Centrifugation of the methylene chlo-
ride can be used if necessary to break emulsions. Phenolic compounds must
be in an aqueous solution to be quantified by the 4-aminoantipyrine pro-
cedure. To meet this requirement, the phenolic compounds are extracted from
the methylene chloride into 0.005M sodium hydroxide solution. The details
of this procedure are presented in Appendix D.
Experiments were performed to determine the optimal conditions for
extraction of phenolic compounds from methylene chloride into a dilute
sodium hydroxide solution. A methylene chloride extract of sludge was
prepared by stirring 50 g of sludge with 1 liter of methylene chloride for
2 hours. The solid particles were filtered from the mixture, and seven
replicate 100-mL aliquots of methylene chloride were spiked with one of each
of the radiolabeled phenolic compounds to give a concentration of 11 pg/L of
phenol, 11 Mg/L of g-nitrophenol, or 130 pg/L of 2,4,5-trichlorophenol.
Acid-soluble compounds were extracted from the methylene chloride with 15 mL
of nitric acid solution and discarded. The phenolic compounds were serially
extracted from the methylene chloride with 20, 10, and 10 mL aliquots of
0.005M sodium hydroxide solution. The sodium hydroxide extracts were com-
bined and diluted to 50 mL with 0.005M sodium hydroxide solution. A control
consisting of clean methylene chloride was spiked and extracted with each
set of sludge-methylene chloride extracts. For all samples, the percentages
of the phenolic compounds present in each extract was determined by liquid
scintillation counting and are presented in Table 15. The average percent-
ages of phenol, nitrophenol, and trichlorophenol recovered from seven repli-
cates are 74, 89, and 86, respectively. Recovery from the controls was
similar, indicating that substances present in the sludge which are soluble
in methylene chloride do not reduce the extraction efficiency of the three
phenolic compounds. In the final form of the method, the cleanup extraction
with nitric acid is not necessary, as the acid-soluble compounds remain in
the sulfuric acid-solid mixture and are not extracted into the methylene
chloride.
The concentration of total phenolic compounds in the sodium hydroxide
solution is determined by the 4-aminoantipyrine procedure. To obtain the
clear sodium hydroxide solution required for the 4-aminoantipyrine proced-
ure, it may be necessary to filter the methylene chloride extract of some
matrices. Loss of phenol from the methylene chloride by filtration was
investigated. Seven replicate solutions of methylene chloride containing
21 pg/L of radiolabeled phenol were filtered through qualitative filter
paper. Aliquots of each sample were subjected to liquid scintillation
counting before and after filtration. The average recovery of phenol in the
seven replicate filtered samples was 95+3 percent.
-------
TABLE 15. AVERAGE PERCENTAGES AND STANDARD DEVIATIONS:OF PHENOL, NITROPHENOL, AND
TRICHLOROPHENOL EXTRACTED FROM METHYLENE CHLORIDE-SLUDGE EXTRACT !
Sodium hydroxide solution t
Nitric
First
Second
Third
1
I
Compound
acid extract
extraction
extraction
extraction
total
phenol
1
sludge
3 + 0
45 + 3
19+1
11 + 1
j 74 + 5
aqueous control
3 + 0
46 + 2
16+2
10 + 1
! 72 + 5
p-nitrophenol
sludge
2
82
3
0.5
i 89 + 3
aqueous control
2
83
2
0.3
i 85
2,4,5-trichlorophenol
•
sludge
3
86
2
0.3
I 86 + 3
aqueous control.
3-
86
2 i
0.3
! 86
*Number of replicate samples.
-------
The. literature was reviewed for information on compounds which inter-
fere with determination of total phenolic compounds by the 4-aminoantipyrine
procedure. Groups of chemicals which are potential interferences are alipha-
tic aldehydes, reduced sulfur compounds, aromatic amines, and certain metals
(108,110). At the pH range of 9.8 to 10.2 at which the reaction is
carried out, the tendency of interfering keto-enol systems to respond to the
.-'procedure is greatly reduced (108). Host metal and sulfur compounds will
not be present in the final methylene chloride extract. Solid and semisolid
environmental samples are not expected to contain any interferences which
have not been found and investigated in aqueous environmental samples.
The reaction of phenolic compounds with 4-aminoantipyrine in the pre-
sence of potassium ferricyanide occurs over a wide pH range. It is impor-
tant, however, to adjust the pH of the solution to the narrow range of
10 ± 0.2 to insure reproducibility, maximum sensitivity, and minimal re-
sponse to interferences. The sodium hydroxide solution extract of the
methylene chloride has a pH of less than 10. The 4-aminoantipyrine proce-
dure, as recommended by the U.S. EPA (9), specifies the addition of ammonium
chloride buffer solution to adjust the pH to 10 ± 0.2. Experiments per-
formed with the sodium hydroxide solution showed that the pH tended to be
about 10.5 after addition of the ammonium chloride buffer solution. This
problem was remedied by adjusting the pH of the sodium hydroxide solution
to 4 by adding a few drops of phosphoric acid solution prior to adding the
ammonium chloride buffer solution. This procedure assures a final pH of
10 ± 0.2.
Details of the method for analysis of solid and semisolid environmental
samples for total phenolic compounds are presented in Appendix D. Estimates
of the precision, accuracy, and sensitivity of the method were calculated
from the results of analysis of replicate samples of sludge. Seven repli-
cate 25-g samples of sludge were analyzed for phenol. Three more groups of
seven replicate 25-g portions of sludge were spiked with 2, 11, and 100 pg
of phenol and analyzed. The direct colorimetric 4-aminoantipyrine procedure
was used for quantification of samples containing more than 50 pg of phenol.
Chloroform extraction of the colored product was used for measurement of
phenol levels less than 50 pg. The micrograms of phenol recovered from each
sample are listed in Table 16. The average recoveries of phenol from sam-
ples spiked with 2, 11, and 100 |jg of phenol were 56, 41, and 45 percent,
respectively. The standard drviations of samples containing 0.7, 2.7, 11.7,
and 100.7 (Jg of phenol were 0.3, 0.8, 0.8, and 4, respectively. The coeffi-
cients of variation, calculated by dividing the standard deviation by the
mean level of phenol recovered, were 43, 53, 17, and 9 percent, respective-
ly, for samples containing 0.7, 2.7, 11.7, and 100.7 pg of phenol. The
method is sensitive to approximately 1 pg of phenol. When a 25-g sample is
extracted, phenol concentrations as low as 0.04 pg/g can be determined.
Further research is needed to improve the recovery of the method.
PHASE IV: VALIDATION OF THE METHOD
The method developed for the analysis of solid and semisolid environ-
mental samples for total phenolic compounds was validated in Phase IV. The
.50
-------
TABLE 16. MICROGRAMS OF PHENOL RECOVERED FROM SPIKED AND UNSPIKED
SAMPLES OF SLUDGE
Replicate
Hg of phenol
added to sample
number
0
2
11
100
1
.1.3
-2.0 ..
.4.2.
-7
2
0.9
0.4
4.0
40
3
0.9
0.6
5.0
50
4
0.6
2.5
6.3
49
5
0.7
1.6
4.8
45
6
0.4
2.3
4.4
42
7
0.4
1.3
4.7
46
average pg recovered
0.7
1.5
4.8
45 .
standard deviation
0.3
0.8
0.8
4
coefficient of
variation
43%
53%
17%
9%
actual |jg of
phenol in sample
0.7
2.7
11.7
100.7
percent of phenol
56%
41%
45%
recovered j
I method consists of solvent extraction of phenolic compounds with methylene
• chloride from an acidified slurry of the solid sample. The phenolic com-
' pounds are extracted from methylene chloride into dilute sodium hydroxide
solution and quantified by the 4-aminoantipyrine colorimetric procedure.
' The method is presented in Appendix D.
i The method for analysis of solid and semisolid environmental samples
• for total phenolic compounds was validated by analysis of ten sample ma-
: trices:
i i
' 1. lake bottom sediment j
2. industrial sludge ]
i 3. industrial solid wastes from a disposal site i
4. fish tissue
j 5. microinvertebrates (shrimp) !
j 6. algae j
! 7. soil contaminated by chemicals from the disposal of industrial
sludges
I 8. growing vegetation —i
=£
r~ r~. . .. ¦' ...
51
-------
9. industrial processing slags disposed of in a landfill
10. commercial incinerator fly ash disposed of in a landfill
One sample of each matrix was obtained, and six 25-g portions of each
sample were frozen to preserve the phenolic compounds. Two portions of each
sample matrix were removed from the freezer and analyzed to determine the
concentration of total phenolic compounds in the samples. The four remain-
ing portions of each matrix were removed from the freezer, thawed, spiked
with an aqueous solution of phenol, held for 16 hours to permit adsorption
of phenol to the matrix, and analyzed for total phenolic compounds. Two
25-g portions of each matrix were spiked with 20 |ig of phenol, and two were
spiked with 100 |jg of phenol to give concentrations of 0.80 and 4.0 IJg/g,
respectively.
The concentrations of phenol recovered from the samples are presented
in Table 17. The recovery of phenol ranged from 22 to 100 percent with an
average of 50 ± 20 percent. .Over the concentration range evaluated, recov-
ery was not dependent on concentration of phenol. An average of 50 ± 25
percent of the phenol was recovered from samples spiked with 20 fJg, and
50 ± 13 percent was recovered from samples spiked with 100 |Jg. Recovery
does appear to be related to the sample matrix. The average percentage of
phenol recovered from the six inorganic matrices—bottom sediment, indus-
trial sludge, industrial solid waste, industrial processing slag, incin-
erator fly ash, and contaminated soil—was 60 ± 19. An average of 33 ± 8
percent was recovered from the organic matrices which were fish tissue,
microinvertebrates, algae, and vegetation.
Quality control samples were extracted and analyzed with each group of
environmental samples. One out of every five samples analyzed was a quality
control sample consisting of distilled water spiked with phenol. The per-
centage of phenol recovered from these samples is presented in Table 18. A
calibration curve was prepared each day using phenol standards prepared in a
0.005M sodium hydroxide solution. In addition, a reagent-glassware blank
was carried through the extraction and quantification procedure daily.
52
-------
TABLE 17. CONCENTRATIONS OF PHENOL ADDED TO AND RECOVERED
FROM SOLID AND SEMISOLID ENVIRONMENTAL SAMPLES
; Average
percent
recovery +
,2; ¦ Concentration Concentration • Percent standard
Matrix added (|Jg/g) recovered (|Jg/g) recovery deviation
! bottom sediments
0
ND
75+ 5
1
0
J ND
' ' '
0.80
! 0.64
80
I
0.80
0.62
78
i
4.0
2.8
70
i
4.0
2.9
72
j industrial sludge
0
ND
55+4
j
0
ND
_ _ •
0.80 . .
0.48
60
!
0.80
0.46
57
4.0
2.1
53
4.0
2.0
50
industrial solid
0
ND
71 + 13
waste
0
0.05
i
: t
0.80
0.56
70
; i
0.80
0.72
90
' !
4.0
2.5
62
t !
: >
4.0
2.5
62
1
j industrial processing
0
81 + 14
i slag
0
i
t '
0.80
0.80
100
'
0.80
0.68
85
•
4.0
2.9
72
:
4.0
2.7
68
. incinerator fly ash
0
: 0.20
42+2
, -
0
. 0.16
j
0.80
0.52
42
i j
0.80
0.48
38
i 1
f !
4.0
. 1.9
43
4.0
! 1.9
43
! contaminated soil
0
ND
36 + 10
!
0
0.08
{
0.80
, 0.24
25
!
i •
0.80
0.28
30
i
4.0
^ 1.8
43
I
4.0
1.8
45
L..
Continued
• ,
. ¦ J
J. •
.53
r it
-------
Table 17 (Continued).
Average
,
percent
i
1
recovery +
Concentration
Concentration
Percent
standard
Matrix
added (^g/g)
recovered (|Jg/g)
recovery
deviation
fish tissue
0
: 0.12
26+3
; 0
; 0.08
0.8
; 0.32
28
0.8
0.28
22
1 4.0
1.2
28
4.0
1.1
26
microinvertebrates 0
0.28
39+5
(shrimp)
0
0.32
0.80
0.64
38
. 0.80
0.56
. 32 .
— ...
4.0
2.1
44
4.0
2.0
42
algae
0
0.20
34 + 10
o
0.12
!" 0.80
0.36
25
0.80
0.36
25
4.0
; 1-9
43
: 4.0
1.8
42
vegetation
0
: 0.16
42+9
0
: 0.16
0.80
: 0.40
30
! 0.80
0.48
40
4.0
1.9
48
4.0
1.9
48
. ND = not detected at concentration above detection limit of 0.05 Mg/g
54
-------
TABLE 18. RECOVERY OF PHENOL FROM AQUEOUS QUALITY ASSURANCE SAMPLES
Sample pg of phenol in sample |Jg of phenol recovered Percent
1.
20
14 -
70
2
20
14
70
3
100
80
80
4
100
i 72
72
5
20
16
80
6
20
14
70
7
100
74
74
8
100
72
72
9
20
14
70
10
20
16
80
11
20
16
80
12
20
16
80
13
100
68
68
14
100
66
66
average ± standard deviation 74 ± 5
i 55
-------
SECTION 5
PRESERVATION OF CYANIDE AND TOTAL PHENOLIC COMPOUNDS IN SOLID AND SEMISOLID
ENVIRONMENTAL SAMPLES
INTRODUCTION
The analysis of environmental samples for cyanide and phenolic com-
pounds at trace levels requires methods which are sensitive, accurate, and
precise. Therefore, carefully controlled laboratory conditions ere essen-
tial. To meet these requirements it is generally necessary to ship samples,
collected in the field, to a centralized laboratory for analysis. This
shipping process can lead to substantial delays between collection and
analysis. The effectiveness of sample preservation techniques can substan-
tially affect the accuracy of data generated by the analysis.
The major processes which adversely affect sample integrity are evapo-
ration of volatile compounds, adsorption of more polar compounds on the
walls of the sample container, and chemical and/or biological degradation of
the sample components. It is possible to slow or eliminate these processes
by adding certain chemicals which will not interfere with the analysis to
the sample at the time of collection and by refrigerating the sample between
the time of collection and analysis (147,148). Unfortunately, a review of
the literature revealed a paucity of information on the preservation of
samples for analysis for cyanide and phenolic compounds. The few mentions
of the employment of preservation techniques for these analyses are re-
stricted to aqueous samples.
Preliminary studies to determine optimal conditions for storage of
samples containing cyanide and phenolic compounds were performed. Solid or
aqueous samples were spiked with radiolabeled cyanide or phenol, and the
concentrations were monitored for 16 days by liquid scintillation cousk ng.
Use of radiolabeled compounds allowed losses by volatilization and adsorp-
tion to be detectec.. Loss by degradation of the compounds could not be
assessed unless the radiolabeled degradation products were lost from the
sample. Three sets of btorage conditions were evaluated for preservation of
aqueous solutions containing cyanide: storage at less than'0°C, storage at
4-6°C, and addition of sodium hydroxide with storage at 4-6°C. Conditions
evaluated for storage of solid samples containing phenol were similar:
storage at less than 0°C, storage at 4-6°C, and addition of phosphoric acid
and copper sulfate followed by storage at 4-6°C. The most favorable condi-
tions for preservation of samples containing cyanide or phenol appeared to
be storage at temperatures of less than 0°C.
Studies were performed to confirm the feasibility of preservation by
-freezing. Samples of a solid matrix were spiked with cyanide and phenol and
-------
•\? ¦ . I
- analyzed by the methods in Appendices B and D. The samples used for valida-
j tion of the analytical methods for cyanide and phenolic compounds were
; frozen for preservation. i
! . I
METHODS OF PRESERVATION '
\
'2" To prevent degradation of sample components during shipment from.the
4 field to the laboratory, samples are usually cooled or frozen. Reduced
1 temperatures slow most chemical reactions and decrease biological activity.
' To eliminate all biological activity, mercuric chloride (149-151) or copper
I sulfate (9,13) can be added to the sample in the field. It has also been
{ suggested that extremes of pH will halt biological degradation as only a few
microorganisms can grow at pH values less than 2 or greater than 10 (152).
These techniques can be incorporated into procedures designed to prevent
other types of sample decomposition as discussed below. j
1
Cyanide
1 !
! The most common procedure for preserving samples for cyanide analysis
, is to acM base in the field to raise the pH of the sample to 12 or above
' (7,9,13). This procedure prevents the formation of the volatile species,
' hydrocyanic acid. This compound is a weak acid (pK of about 10). Con-
: ditions of high pH prevent protonation of the cyanide ion and thus keep this
j species in solution. Oxidizing agents such as chlorine which will decompose
¦ cyanide are reduced by adding ascorbic acid prior to pH adjustment (9).
j Sulfide in the sample will convert cyanide ion to thiocyanate if it is not
I removed by precipitation with cadmium before preservation. Because cyanides
| are unstable, analysis within 24 hours of collection is recommended (9,13).
f
, Phenolic Compounds
; Generally, samples collected for analysis for total phenolic compounds
j are acidified in the field and shipped refrigerated to the laboratory
! (9,13,149). Copper sulfate can be added to the sample at the level of
j 1 g/L to inhibit biological degradation (9,13,106). Lowering the pH of the
' sample to approximately 4 insures the complete protonation of the hydroxyl
. group of even the most polar substituted phenols. Phenol itself has a
; pK of approximately 10, while the pK of trichlorophenol is 6. Protonated
i phenols are less reactive and less polar than the unprotonated form of these
| compounds. Therefore, the processes of chemical degradation and adsorption
\ onto container walls are slowed or eliminated by acidifying the sample,
j Nevertheless, it is recommended that samples be analyzed within 24 hours of.
i collection.
I
j EVALUATION OF PRESERVATION METHODS
|
i Samples were spiked with radiolabeled cyanide and phenol to determine
| optimal preservation conditions. The change in concentration over time was
] monitored by liquid scintillation counting. The use of radiolabeled cyanide
!
t ' ^ ; ~
"57
-------
and phenol made it possible to detect losses by volatilization and adsorp-
tion. Loss by degradation could not be assessed by liquid scinti_lation
counting.
Cyanide
A method for achieving reproducible, high recoveries of cyanide from
solid samples had not been developed at the time the preservation study was
initiated. For this reason, the preservation conditions were evaluated for
samples of water containing cyanide. Semisolid matrices were used for the
validation experiments. A slightly basic solution containing 50 |Jg/L of
radiolabeled cyanide as potassium cyanide was prepared. Five-milliliter
aliquots of this solution were placed in 15-mL glass vials. The vials were
stored under three sets of Conditions. One set was stored at 4-6°C and
another set at less than 0°C. The third set was treated with 0.5 mL of
2M sodium hydroxide solution to raise the pH to 12 end stored at 4-6°C.
Four samples from each storage group were analyzed by liquid scintil-
lation counting on days 0, 1, 2, 3, 6, 8, 13, and 16. The average percent
recoveries of radiolabeled cyanide in aqueous samples under each storage
condition are summarized in Table 19. Between day 0 and day 1, an average
of 19 percent of the cyanide was lost from all three storage groups. The
percentage of cyanide recovered from the samples stored at 4-6°C decreased
an additional 14 percent between the first and second days, whereas recovery
from the samples stored under the other two conditions decreased only 2 to
| 3 percent. No further significant loss of cyanide occurred until day 13,
i when a mean loss of 17 percent was observed in samples stored under all
conditions.
I
TABLE 19. PERCENT RECOVERY OF UC-LABELED CYANIDE FROM
WATER SAMPLES STORED UNDER VHREE DIFFERENT CONDITIONS
Days of storage
Storage conditions
0
1
2
3
6
8
13
16
<0°C
102
82
80 |
74
75
75
60
58
4°-6°C
101
80
i
66
61
54
54
37
29
4°-6°C, NaOH
96
\
80
i
77
79
80
82
63
62
Phenol ^
Preservation studies to determine optimal ^storage conditions and hold-
ing time for solid samples containing phenol were conducted in Phase II us-
58.
-------
j ing samples of sludge and distilled water. Five-gram portions of wet,
j domestic organic sludge were placed in glass bottles, spiked with 1.8 pg of
;¦ radiolabeled phenol, and stored under one of three sets of conditions.
! Five-milliliter aliquots of distilled water in glass bottles were spiked
i with 0.4 pg of radiolabeled phenol and stored under the same conditions as
:¦ the sludge samples. One set of water and sludge samples was stored at -7°C,
¦and a second set was stored at 6°C. The third set of samples was treated
< with 0.5 mL of concentrated phosphoric acid and 0.5 mL of O.AM cupric sui-
; fate solution and stored at 6°C.
J
I
: Four samples of sludge from each storage group were extracted with
! methylene chloride by the stir-bar extraction procedure on days 0, 1, 2,
1 5, 8, and 16. Radiolabeled phenol in aliquots of the extract was counted to
' determine the recovery of phenol from the sample. Each day on which sludge
! samples were extracted, four additional fresh sludge samples were spiked
with radiolabeled phenol and extracted. These samples served as controls
and permitted monitoring of the extraction efficiency on each day. Aliquots
of the water samples were taken directly from the sample bottles and count-
ed.. Controls for the water samples were prepared each day by spiking 5 mL
of water in a glass bottle with radiolabeled phenol. An aliquot of water
from each control was removed for liquid scintillation counting.
The percentages of phenol recovered from samples of sludge and water in
each storage group are shown in Table 20. No significant loss occurred from
the sludge samples stored at -7°C. The percentage of phenol recovered from
the samples stored at 6°C and those treated with acid and cc >er sulfate and
j stored at 6°C dropped rapidly to 5 percent by the fifth day. Recovery of
j phenol from the controls was near 100 percent each day, except day 0 for
i sludge, indicating that there was no loss of phenol during extraction and
! handling of samples.
i
Conclusions
I
; Of the conditions evaluated for preservation of cyanide in water, the
; most favorable were storage at temperatures of less than 0°C and treatment
1 with sodium hydroxide solution preceding storage at 4-6°C. It is not feas-
; ible to mix a solid sample with a solution of sodium hydroxide in the field,
i although a sample could be immersed in the solution. Samples for cyanide or
; phenol analysis may be frozen in the field using dry ice.
VALIDATION OF PRESERVATION METHODS
Freezing was tested as a method for preservation of solid and semisolid
environmental samples containing cyanide and phenolic compounds.
Cyanide
j Samples of sludge from a chemical manufacturing waste treatment process
! were analyzed for cyanide by the distillation pyridine-barbituric acid
! method (Appendix B). The samples contained 45 pg/g of cyanide, and there-
j~fore, were good candidates for the preservation study. Five-gram portions
-------
TABLE 20. PERCENT RECOVERY OF UC-LABELED PHENOL FROM SAMPLES OF
SLUDGE AND WATER STORED UNDER THREE DIFFERENT CONDITIONS
Days of storage
. Storage conditions
0
1
2
5
8
16
Sludge
;
-7°C
107
100
93
99
98
94
6°C
99
62
16
55
3
3
6°C, H3P04,CuS04
104
54
17
5
3
2
Control*
84
98
100
103
104
95
Water
1
-7°C
103
104
101
97
101
102
6°C
102
104
103
103
103
99
6°C, H3P04, CuSO^
102
103
101
102
101
99
Control*
102
104
103
106
98
103
^Control samples were prepared and extracted each day.
: i
: of sludge were placed in glass vials and frozen. Four replicate samples
: were analyzed for cyanide on days 0, 1, 2, 4, 8 and 16. The concentrations
' of cyanide measured in th= samples are presented in Table 21. No loss
: occurred over the 16 day period.
TABLE 21. CONCENTRATIONS OF CYANIDE (pg/g) IN SLUDGE SAMPLES
STORED AT TEMPERATURES OF LESS THAN 0°C
Replicate
number
0
1
Days of storage
2 4
8
16
1
45
51
' 41
54
36
48
2
50
49
, 43
56
41
46
3
42
43
! 37
54
35
47
4 ;
44
42
,45
52
38
42
average ±
standard deviation
45±3
46±4
42±3
54±2
38+2
46±3
—r
60
.1
—-..J
-------
Phenol
A sample of soil contaminated with phenolic compounds was obtained from
a chemical manufacturing plant. Replicate 8-g portions of soil were placed
in glass vials and frozen. Four replicate samples were extracted and ana-
lyzed on days 0, 1, 2, 4, 8, and 16 by the procedure presented in
Appendix D. - The results of the study, reported in Table 22, show that there
was no significant loss of phenol over the 16-day period.
I
1
TABLE 22. CONCENTRATIONS OF PHENOL (pg/g) IN SOIL SAMPLES
STORED AT TEMPERATURES OF LESS THAN 0°C
Replicate Days of storage
number
0
1
2
4
8
16
1
181
146
164
165
151
175
2
186
159
173
165
160
162
' 3
191
117*
178
159
177
163
4
149
162
168
143
176
179
average ±
177±19
146±20
171±6
158±10
166±13
170±9
standard deviation
* Sample contained a stone. This probably accounts for the lower concentra-
tion of phenol.
. Conclusions !
Storage of samples at temperatures of less than 0°C appears to be an
' effective means of preserving solid and semisolid environmental samples
which are to be analyzed for cyanide or total phenolic compounds. There was
1 no significant loss of cyanide or phenolic compounds from solid, environ-
mental samples stored for 16 days. Advantages of freezing over other pre-
servation techniques are that acids, bases, or other reagents do not have to
' be taken into the field, sample handling is reduced, and introduction of
contaminants is minimized. Freezing was used to preserve the solid and
semisolid environmental samples analyzed in the validation study.
.61
-------
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I
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APPENDIX A
PROCEDURES FOR QUANTIFICATION OF CYANIDE
DETERMINATION OF CYANIDE: PYRIDINE-BARBITURIC ACID METHOD (9)
Reagents
1.1 Sodium hydroxide solution, I.25M: Dissolve 50 g of NaOH in
distilled water, and dilute to 1 liter with distilled water.
1.2 Sodium dihydrogen phosphate, 1M: Dissolve 138 g of NaH^PO^'H^O in
1 liter of distilled water. Refrigerate this solution.
1.3 Stock cyanide solution: Dissolve 2.51 g of KCN and 2 g of KOH
in 1 liter of distilled water. Standardize with 0.0192M AgNO^.
Dilute to appropriate volume so that 1 mL = 1 mg CN.
1.4 Standard cyanide solution A: Dilute 50 mL of stock (1.3) to
1 liter with distilled water (1 mL = 50 pg CN).
1.5 Standard cyanide solution B: Dilute 100 mL of standard cyanide
solution A to 1 liter with distilled water (1 mL = 5 |Jg CN).
1.6 Chloramine-T solution: ileigh and dissolve 1.0 g of white, water
soluble Chloramine-T in 100 mL of distilled water and refrigerate
until ready to use. Prepare fresh weekly.
1.7 Pyridine-Barfcituric acid reagent: Put 15 g of barbituric acid in
a 250-mL volumetric flask. Add just enough distilled water to
wash the sides of the flask and wet the barbituric acid. Add
75 mL of pyridine and mix. Add 15 mL of concentrated hydrochloric
acid (sp. gr. 1.19), mix and cool to room temperature. Dilute to
250 mL with distilled water and mix. This reagent is stable for
approximately six months if stored in a cool, dark place.
I 1
Procedure i
2.1 Prepare a series of standards by pipetting suitable volumes of
standard cyanide solution B (1.5) into 250 mL volumetric flasks.
To each standard add 50 mL of 1.25M sodium hydroxide solution
(1.1) and dilute to 250 mL with distilled water. Prepare as
follows: \
-------
mL of Standard Solution B pg of cyanide
(1.0 mL = 5 (Jg cyanide) per 250 mL
0 0
1.0 5
2.0 10
5.0 25
10.0 50
15.0 75
20.0 100
2.2 Withdraw 50 mL of each standard and transfer to 100 mL volumetric
flasks. Also withdraw 50 mL of the distilled samples and transfer
to 100 mL volumetric flasks.
2.3 Add 15 mL of sodium phosphate solution (1.2) to all of the
standards and samples and mix.
2.4 Add 2 ml. of Chloramine-T solution (1.6) and mix. Allow to stand
1 to 2 minutes.
2.5 Add 5 mL of pyridine-barbituric acid reagent (1.7) and mix.
Dilute to mark with distilled water and mix again.
2.6 Allow 8 minutes for color development, and then read the absor-
bance at 578 nm in a 1 cm cell within 15 minutes.
Calculations
3.1 Prepare a standard calibration curve by plotting absorbance of
standard versus cyanide concentration in iJg per 250 mL.
3.2 Determine the unknown sample concentration corresponding to the
measured absorbance from the calibration curve.
3.3 Calculate the cyaiiide concentration, in Mg/L, in the original
sample fit. follows:
„ Ax 1,000 50
Cyanide, fJg/L - jjJ x —
where A = |Jg cyanide read from standard curve
B = mL of original sample for distillation
¦ C = mL taken for colorimetric analysis
-------
DETERMINATION OF CYANIDE: TRIS (1,10-PHENANTHR0LINE)-IR0N II
COMPLEXATION (46)
Reagents
1.1 Stock cyanide solution: Dissolve 2.51 g of KCN and 2 g of KOH in
- ---I liter of distilled water. Standardize with 0.0192M AgNO^.
Dilute to appropriate concentration so that 1 mL = 1 mg CN.
1.2 Intermediate cyanide solution: Dilute appropriate volume of the
stock cyanide solution to obtain an intermediate working solution
having CN concentration of 10.0 pg/mL.
1.3 Disodiuin hydrogen phosphate solution, 1M: Dissolve 381.37 g of
Na2HP0^*7H20 and dilute to 1 liter with distilled water.
1.4 Hydroxylamine hydrochloride solution, 10%: Dissolve 100 g of
hydroxylamine hydrochloride (NH^OH'HCl) in 1 liter of distilled
water. r r
1.5 Sodium hydroxide solution, 0.5M: Dissolve 2 g of NaOH in
distilled water and dilute to 100 mL.
1.6 Thymol blue indicator, 0.1%: Weigh 0.1 g of thymol blue indicator
into a 100 mL volumetric flask. Add a small volume of 0.5M NaOH
solution (1.5), swirl to dissolve, and dilute to volume with
distilled water.
1.7 Ferroin Reagent Solution: Dissolve together 1.96 g of Fe(NH^)2 -
(SO^^^H^O and 3.17 g of 1,10-phenanthroline raonohydrate to a
volume of 1 liter using distilled water. The resulting solution
is 0.005M ferroin su]fate and 0.001M 1,10-phenanthroline.
1.8 Acetic acid solution, 1M: Dilute 58 mL of glacial acetic acid to
1 liter with distilled water.
1.9 Chloroform
Procedure
2.1 Prepare a series of standards by diluting appropriate volumes of
the intermediate cyanide solution. Standards should have a CN
concentration range of 0.2 to 1.0 (Jg/mL.
2.2 Measure 25 mL of the standards and the samples into 50 mL glass-
stoppered Erlenmeyer flasks.
2.3 Add 5 mL of 1M disodiuin hydrogen phosphate solution (1.3), 1 mL
of 10% hydroxylamine hydrochloride solution (1.4), and 2 drops of
thymol blue indicator (1.6). If the indicator does not impart a
yellow color to the solution, add 1M acetic acid (1.8) drop-by-
drop until the yellow color persists. - -- - —%
- 76
-------
2.4 Add 0.5M sodium hydroxide solution (1.5) drop-by-drop until the
indicator changes to blue; an intermediate light green color will
be observed just before this point is reached.
2.5 Add 5 mL of the ferroin reagent (1.7) and heat in a boiling water
bath for 10 to 15 minutes with the glass stopper in place.
2.6 Cool the contents of the flasks and transfer to 60-raL separatory
funnels. Extract each solution four times with 5 ml portions of
chloroform. Combine the extracts by filtering into 25-mL volumet
ric flasks and diluting to final volume with chloroform.
2.7 Measure the absorbance of the chloroform extracts at 597 nm in a
1-cm cell using chloroform as the standard blank.
Calculations
3.1 Prepare a calibration curve by plotting the absorbance versus
concentration of the standards.
3.2 Determine the sample concentration corresponding to the measured
absorbance from the calibration curve.
3.3 If the sample was distilled prior to determination, calculate the
concentration of cyanide in the original sample as follows:
where A = |Jg/mL cyanide from curve
B = volume of distillate (mL)
C = volume of original sample taken for distillation (mL)
Hg/mL
cyanide in
sample
A x B
C
77
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DETERMINATION OF CYANIDE: MERCURIC CHLORANILATE - U.V. METHOD (49)
Reagents
1.1 Mercuric chloranilate: In a small beaker, gently wash approxi-
mately 10 to 15 g of mercuric chloranilate powder 3 or 4 times
„ .—using small volumes of ethanol. Filter and spread the washed
, chloranilate in a clean beaker and allow to dry at room tempera-
ture.
1.2 Stock cyanide solution: Dissolve 2.51 g of KCN and 2 g of KOH in
1 liter of distilled water. Standardize with 0.0192M AgNO^.
Dilute to appropriate concentration so that I mL = 1 mg CN.
1.3 Intermediate cyanide solution: Dilute appropriate volume of the
stock cyanide solution to obtain an intermediate working solution
have CN concentration of 10.0 pg/mL.
1.4 Ethanol-water solvent: Mix 300 mL of 95% reagent grade ethanol
and 200 ml of distilled water. f
1.5 Acid Buffer: Dilute 10 mL of concentrated nitric acid to 100 mL
with distilled water.
Procedure
2.1 Prepare a series of standards by pipetting appropriate volumes of
the intermediate cyanide solution (1.3) into 50 mL volumetric
flasks. Dilute to volume with the ethanol-distilled water solvent
(1.4). Prepare as follows:
Volume of 10 (Jg/mL Concentration of
cyanide solution cyanide, ps/mL
0
0
1
mL
0.2
2
mL
0.4
5
mL
1.0
10
mL
2.0
20
mL
4.0
40
mL
6.0
2.2 Prepare samples by transferring an appropriate volume to a 50 mL
volumetric flask and diluting with ethanol-water solvent.
2.3 Withdraw 10-mL aliquots from each standard and sample and place in
centrifuge tubes. Add 0.01 g of the washed mercuric chloranilate
powder (1.1) and cap the tubes. Shake vigorously, allow to stand
for 15 to 20 minutes, and then centrifuge at a moderate speed for
20 minutes.
2.4 Adjust the pH of the centrifuged solutions to 4.0 by drop-by-drop
78
-------
addition of the acid buffer solution (1.5) and gently shake to'
mix. Do not attempt to disturb the excess mercuric chloranilate
that has settled to the bottom of the tube.
2.5 Read the absorbance of the standards and samples at 330 nm in a
1 cm absorption cell.
Calculations
3.1 Plot the absorbance versus the concentration of the standards.
3.2 Determine the sample concentration corresponding to the measured
absorbance from the calibration curve.
3.3 If the sample was distilled prior to determination, calculate the
concentration of cyanide in the original sample as follows:
°f A x B
cyanide in =
sample C
where A = (Jg/mL of cyanide from curve
B = volume of distillate (mL)
C = volume of original sample taken for distillation (mL)
.79
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DETERMINATION OF CYANIDE: AMMONIACAL NiCl2 METHOD (5)
Reagents
1.1 Stock cyanide solution: Dissolve 2.51 g of KCN and 2.0 g of KOH
in 1 liter of distilled water. Standardize with 0.0192M AgNO^.
... ... Dilute to appropriate concentration so that 1 mL = 1 mg CN.
1.2 Intermediate working solution: Dilute appropriate volumes of the
stock cyanide solution to obtain intermediate working solution
having a CN concentration of 10 pg/mL.
1.3 Potassium hydroxide solution, 0.25M: Dissolve 9 g of potassium
hydroxide in distilled water and dilute to 1 liter.
1.4 Ammoniacal nickel chloride solution: Dissolve 0.2268 g of nickel
chloride in 500 mL of distilled water in a 1-liter volumetric
flask. Slowly add 67 mL of concentrated ammonium hydroxide.
Wash sides of volumetric flask with distilled water, mix, and di-
lute to the mark. This solution is 1M in NH^ and 0.002M in NiCl^
Procedure
2.1 Prepare a series of standards by pipetting suitable volumes of the
intermediate cyanide working solution (1.2) into 50 mL volumetric
flasks and diluting to volume with the 0.25M KOH solution (1.3).
Prepare as follows:
Volume of 10 (Jg/mL
cyanide solution
0
1 raL
2 mL
5 mL
10 mL
20 mL
30 mL
Concentration of
cyanide, Mg/mL
0
0.2
0.4
1.0
2.0
4.0
6.0
2.2 Into test tubes or small vials, measure 5 mL of standards and
samples. (
* \
2.3 Add 5 mL of the ammoniacal nickel chloride solution (1.4) and
shake to mix. Let stand for 20 minutes to allow the reaction to
occur.
\
2.4 Read absorbance at 267 nm.
\
Calculations
3.1 Plot absorbance versus concentration of the^standards.
80
-------
3.2 Read the sample absorbance off the calibration curve to obtain the
concentration of the sample.
3.3 If the sample was distilled prior to determination, determine the
-concentration of cyanide in the original sample as follows:
. A x B
cyanide in = —-—
sample
where A = Hg/mL cyanide from curve
B = volume of distillate (mL)
C = volume of original sample taken for distillation (mL)
81
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DETERMINATION OF CYANIDE: REACTION OF CYANIDE WITH
ORGANIC DISULFIDES USING DIMETHYLFORMAMIDE (51)
Reagents
1.1 Stock cyanide solution: Dissolve 2.51 g of KCN and 2 g of KOH in
-liter of distilled water. Standardize with 0.0192M AgNO^.
Dilute to appropriate concentration so that 1 mL = 1 mg GN.
1.2 Intermediate cyanide solutions: Make appropriate dilutions from
the stock CN solLtion to obtain intermediate working solutions
having CN concentrations of 30 (Jg/mL and 3.0 pg/mL.
1.3 pH 7 buffer solution: Dissolve 3.549 g of Na„HP0,*7H„0 and
3.403 g KH„P0, in 1 liter of distilled water. Dilute to the mark.
2 4
1.4 pH 9 buffer: Weigh and dissolve 3.8137 g of Na^B^O^'lOH^O and
dilute to 1 liter with distilled water.
1.5 DTNB solution, 0.001M: Carefully weigh 0.09908 g of 5,5'-dithio-
bis 2-nitrobenzoic acid. Dissolve in 25 mL of 95% reagent grade
ethanol and dilute to 250 mL with the pH 7 buffer solution (1.3).
1.6 DMF, N,N-dimethylformamide: May be obtained commercially.
Procedure
2.1
Prepare a series of standards by transferring appropriate volumes
of the intermediate cyanide working solutions (1.2) into 50 mL
volumetric flasks. Prepare as follows:
Cyanide
Concentration (pg/mL)
Volume (mL)
2
5
,1
2
3
4
5
Intermediate
cyanide solution (pg/mL)
9 «nn<
3 ppm
3 ppm
30 ppm
30 ppm
30 ppm
30 ppm
30 ppm
0.12
0.3
0.6
1.2
1.8
2.4
3.0
2.2 Pipette from 0.5 to 5 mL of sample into 50 mL volumetric flasks.
2.3 To the 50 mL volumetric flasks containing the cyanide standards
and samples, add 10 mL of pH 9 buffer (1.4), and mix. Add 10 mL
of DMF (1.6) and mix. Add 5 mL of DTNB solution (1.5), nix, and
dilute to 50 mL with the pH 9 buffer (1.4).
2.4 Read the absorbance at 325 nm within 30 minutes after dilution,
using the pH 9 buffer to adjust the spectrophotometer reading to'
zero.
-------
i^v-
Calculations
3.1 Prepare a calibration curve by. plotting the absorbance versus
concentration of the standards.
i
3.2 Determine the unknown sample concentration corresponding to the
measured absorbance from the calibration curve. .
3.3 If the original sample was distilled prior to determination,
calculate the concentration of cyanide in the original as follows:
(jg/mL of
cyanide in
sample
A x B
i-.r •
where A = |Jg/mL cyanide from curve
B = volume of distillate (mL)
,C = volume of original sample taken for distillation (ml)
I .
I t
;i o,"\'
r-
6HA-207 (Cin.i
(1-V6)
-------
DETERMINATION OF CYANIDE: ION SELECTIVE ELECTRODE
1;.. Reagents
1.1 Stock cyanide solution: Dissolve 2.51 g of KCN and 2 g of KOH in
1 liter of distilled water. Standardize with 0.G192M AgNO^.
Dilute to appropriate concentration so that 1 mL = 1 mg CN.
Intermediate working solutions: Dilute appropriate volumes of the
stock cyanide solution to obtain intermediate working solutions
having CN concentrations of 100 pg/mL, 10 pg/mL, and 1.0 pg/mL.
' . I
10M NaOK solution: Dissolve 400 g of NaOH in distilled water and
dilute to 1 liter with distilled water. This is the ionic
strength adjustor (ISA).
2. Apparatus
2.1 Cyanide ion selective electrode
2.2 Reference electrode
2.3 pH/rav meter
2.4 Magnetic stirrer
2.5 Stir bars
3. Procedure
' 3.1 Calibration of electrode
3.1a Put 100 mL of distilled water and 1 mL of ISA solution into a
,250-mL beaker. Place the el-;Erodes in the solution to a
depth of about 3 cm. Begin stirring and set the meter to
read millivolts.
3.1b Pipet 1 mL of the stock cyanide solution into the solution.
Stir thoroughly. Read electrode potential in millivolts and
record.
3.1c Add 10 mL of stock CN solution. Stir thoroughly. Read
electrode potential in millivolts and record.
3.Id Determine the difference between the first and second potent-
ial readings. Correct electrode operation is indicated by a
difference of 58-59 mV, assuming the solution temperature is
between 20°C and 25°C.
3.2 Direct measurement
1.2
1.3
-------
TVHiAir-J Gl »fIjE SH'£)£T
3.2a ,Measure 100 mL of the 10 pg/mL cyanide working solution into"
'a 250 ml beaker and add 1 mL of ISA solution. Set the rela-
tive mV to 000.0 using the 10 pg/mL standard. Read the elec-
itrode potential of the 10 (ig/mL solution.
3.2b Rinse electrodes, blot dry, and place in 100 mL of the
1 Mg/mL cyanide working solution. Add 1 mL of ISA, stir
thoroughly, and read the electrode potential. Record the most
stable reading.
J
3.2c Rinse electrodes, blot dry, and place in 100 mL of the
100 |Jg/mL cyanide working solution. Add 1 mL of ISA, stir
thoroughly, and read the electrode potential.
3.2d Rinse electrodes, blot dry, and place in 100 mL of sample.
Add 1 mL ISA. Stir thoroughly. Record the most stable
^millivolt reading.
3.-3 Low level measurement
3.3a Add 100 mL of distilled water and 1 mL of ISA to a beaker.
Place electrodes in this solution. Stir thoroughly.
3.3b Add increments of the 10 pg/mL cyanide solution using the
table outlined below. Measure the electrode potential after
;each addition.
Added Resulting
volume of concentration
10 mg/L cyanide mg/L
0.1 mL 0.01
0.1 mL 0.02
0.2 mL 0.04
0.2 mL 0.06
0.4 mL 0.10
2.0 mL 0.29
2.0 mL 0.48
3.3c Rinse electrodes, blot dry, and place in 100 mL of sample.
Add 1 mL of ISA solution. Stir thoroughly. Wait for a
[stable potential reading and record.
i
4. Calculations
4.1 Direct measurement
4.1a Plot the millivolt readings (linea:: axis) against concentra-
tion (log axis) on semi-logarithmic graph paper.
4.1b Determine the sample concentration from the calibration
•curve.
85
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TViWCj GtMOE SMFO
4.1c .If the samples have been distilled prior to determination,
calculate the cyanide concentration in the original sample as
follows: t
|jg/mL of A x B
cyanide in = C
sample
where A - |Jg/mL cyanide from curve
B = volume of distillate (mL)
C = volume of original sample taken for distillation (mL)
4.2 Low-level measurement
¦ 4.2a Plot the millivolt readings (linear axis) against concentra-
tion (log axis) on semilogaritlunic paper.
4.2b Determine the sample concentration corresponding to the
measured, potential from the low-level calibration curve.
4.2c If the samples have been distilled prior to determination.,
calculate the cyanide concentration in the original sample
follows:
pg/mL of A x B
cyanide in - C
sample
where A = (Jg/mL cyanide from curve
B = volume of distillate (mL)
C = volume of original sample taken for distillation (mL)
;.8
-------
-TYP'iPJG GUiDr'SHiTT
APPENDIX B
METHOD FOR ANALYSIS OF SOLID AND SEMISOLID ENVIRONMENTAL SAMPLES FOR CYANIDE
CYANIDE, TOTAL
1. Scope and Application
1.1 This method is applicable to the determination of cyanide in solid
and semisolid environmental samples.
1.2 The procedure can be used for determination of cyanide concentra-
tions exceeding 0.5mg/g in a 10-g sample.
2. Summary of Method
2.1 Cyanide as hydrocyanic acid (HCN) is released from cyanide
complexes by means of a reflux-distillation operation and absorbed
in a sodium hydroxide solution. The cyanide ion in the absorbing
solution is determined colorimetricaliy.
2.2 In the colorimetric determination, cyanogen chloride, formed by
reaction of cyanide with chloramine-T at a pH less than 8, reacts
with barbituric acid in a solution of pyridine to form a colored
product. The absorbance is measured at 578 run.
3. Definitions
3.1 Cyanide is defined as cyanide ion and complex cyanides converted
to hydrocyanic acid by reaction in a reflux system of a mineral
acid in the presence of magnesium ion.
A. Sample Handling and Preservation
4.1 The sample should be collected in plastic or glass wide-mouth
bottles. Sample bottles should be thoroughly cleansed and rinsed
to remove soluble material.
4.2 Samples should be frozen as soon as possible after col lection.
Samples can be held for up to 16 days without significant los3 of
cyanide.
5. Interferences
5.1 Interferences are reduced or eliminated by using the distillation
procedure. Volatile sulfur-containing compounds are removed
87
-------
during distillation by bubbling the distillate through a lead
acetate solution.
Apparatus
6.1 Reflux distillation apparatus such as that shown in Figure 1. The
boiling - flask should be 1 liter in size and equipped with an air
inlet tube and condenser. The sulfur-removal vessel should be
equipped with an open-ended glass tube, and the cyanide absorber
tube should be equipped with a glass tube with fritted end.
6.2 Spectrophotometer suitable for measurement at 578 run with a
1.0 cm or larger cell.
6.3 Volumetric flasks, 25G ml, 100 ml.
Reagents
7.1 Sodium hydroxide solution, 1.8M: Dissolve 72 g of NaOH in
distilled water and dilute to 1 liter with distilled water.
7.2 Dilute sodium hydroxide solution, 0.25M: Dilute 140 mL of sodium
hydroxide solution (7.1) to 1 liter with distilled water.
7.3 Sulfuric acid, concentrated.
7.4 Magnesium chloride solution: Dissolve 510 g of MgCl2'6H20 in
distilled water and dilute to 1 liter with distilled water.
7.5 Lead acetate solution: Dissolve 30 g of Pb(C H Oj) '3H20 i*i
950 mL of distilled water. Adjust the pH to £.5 with acetic acid
and dilute to 1 liter with distilled water.
7.6 Sodium dihydrogenphosphate, 1M: Dissolve 138 g of NaHjPO^-H^O in
1 liter of distilled water. Refrigerate this solution.
7.7 Stock cyanide solution: Dissolve 2.51 g of KCN and 2 g of KOH
in 1 liter of distilled water. Standardize with 0.0192M AgN0_.
Dilute to appropriate volume so that 1 mL = 1 mg CN.
7.8 Standard cyanide solution, intermediate: Dilute 50.0 mL of stock
(1 mL = 1 mg CN) to 1000 mL with distilled water (1 mL = 50.0 mg).
7.9 Standard cyanide working solution: Prepare fresh daily by dilut-
ing 100 mL of intermediate cyanide solution to 1000 mL with dis-
tilled water and store in a glass stoppered bottle.
1 mL = 5.0 mg CN.
7.10 Standard silver nitrate solution, 0.0192M: Prepare by crushing
5 g of AgNO^ crystals and drying to a constant weight at 40°C.
Weigh ouc 3.2647 g of dried AgNO , dissolve in distilled water,
and dilute to 1000 mL (lmL =1 mg CN).
88
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VPiiNiG OLJiOC SHifri
7.11 Chloramine-T solution: Dissolve 1.0 g of white, water soluble
chloram.!.ne-T in 100 mL of distilled water and refrigerate until
ready to use. Prepare fresh weekly.
7.12 Pyridine-barbituric acid reagent: Place 15 g of barbituric acid
in a 250 mL volumetric flask and add just enough distilled water
to wash the sides of the flask and wet the barbituric acid. Add
75 mL of pyridine and mix. Add 15 mL of HC1 (sp gr 1.19), mix,
and cool to room temperature. Dilute to 250 mL with distilled
water. This reagent is stable for approximately six months if
stored ir» a cool dark place.
Procedure
8.1 Samples consisting of large pieces of solid material such as
tissue should be chopped in a blender or ground with a grinder.
Place 10 g of wet homogenized sample and 500 mL of distilled water
in a 1 liter boiling flask.
8.2' Add 35 mL of sodium hydroxide solution (7.1) to the cyanide absor-
ber tube and dilute if necessary with distilled water to obtain
an adequate depth of liquid in the tube. Place 25 mL of lead ace-
tate solution in the sulfur removal vessel. Connect the boiling
flask, condenser, absorber, and trap in the train.
8.3 Start a slow flow of air entering the boiling flask by adjusting
the vacuum source. Adjust the vacuum so that there is approxi-
mately one inch of foam on the surface of the sodium hydroxide
solution in the absorber tube. Caution: The air flow rate will
not remain constant after the reagents have been added and while
heat is being applied to the flask. Adjust the air flow rate as
necessary to prevent the solution in the boiling flask from back-
ing up into the air inlet tube.
8.4 Slowly add 25 mL of sulfuric acid (7.3) through the air inlet
tube. Rinse the tube with distilled water and allow the air flow
to mix the flask contents for 3 minutes. Pour 20 mL of magnesium
chloride solution (7.4) into the flask through the air inlet tube
and wash down with distilled water.
8.5 Heat the mixture to boiling, taking care to prevent the mixture
from backing up into the air inlet tube. Reflux for 2 hours.
Turn off the heat but continue the air flow for at least 15 min-
utes. After the boiling flask has cooled, disconnect the absor-
ber and turn off the vacuum source.
8.6 Drain the solution from the absorber into a 250 mL volumetric
flask and bring up to volume with distilled water washings from
the absorber tube.
89
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TYi'l.MG GUIDti SUGEV
8.7 Prepare a series of standards by pipeting suitable volumes of
standard solution into 250 mL volumetric flasks. To each standard
add 35 mL of 1.8M sodium hydroxide solution (7.2) and dilute to
250 mL with distilled water. Prepare as follows:
mL of standard solution pg of CN
- (1-0 mL = 5 MR CN) . per 250 mL
0 0
1.0 5
2.0 10
5.0 25
10.0 50
15.0 75
20.0 100
8.8 Withdraw 50 mL of each standard solution and 50 mL or less of the
sample solution from the flask and transfer to 100 mL volumetric
flasks. If less than 50 mL of a sample solution is taken, dilute
to 50 mL with 0.25M sodium hydroxide solution (7.2). Add 15.0 mL
of sodium phosphate solution (7.6) and mix.
8.8.1 Pyridine-Barbituric Acid Method: Add 2 mL of chlor-
amine-T (7.9) and mix. After 1 to 2 minutes, add 5 mL
of pyridine-barbituric acid reagent (7.11) and mix.
Dilute to mark with distilled water and mix again.
Allow 8 minutes for color development, and read absor-
bance at 578 nm in a 1 cm cell within 15 minutes.
8.8.2 It is not imperative that all standards be distilled in
the same manner as the samples. It is recommended that
at least two standards (a high and low) be distilled and
compared to similar values on the curve to insure that
the distillation technique is reliable. If distilled
standards do not agree closely with the undistilled
standards, the operator should find the cause of the
apparent error before proceeding.
8.8.3 Prepare a standard curve by plotting absorbance of
standard versus |Jg of cyanide.
8.8.4 To check the efficiency of the sample distillation, add
an increment of cyanide from either the intermediate
standard (7.7) or the working standard (7.8) to insure a
level of 20 pg/1 or a significant increase in absorbance
value. Proceed with the analysis as in Procedure 8.1
using the same flask and system from which the previous
sample was just distilled.
90.
-------
TVi'i.^G nsjio? SHftr
9. Calculation
9.1 Calculate the cyanide concentration, in pg/g, in the original
sample as follows:
CN(pg/g) =| x I5
where:
A = (Jg of CN read from standard curve
B = weight of original sample distilled (g)
C = ml taken for colorimetric analysis
91
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"fi?WG GUiDC
APPENDIX C
PROCEDURES FOR QUANTIFICATION OF PHENOLIC COMPOUNDS
DETERMINATION OF PHENOL: 4-AMINOANTIPYRINE METHOD (9)
1. Reagents
1.1 Stock phenol solution: Dissolve 1 g of phenol in 500 mL of dis-
tilled water and dilute to 1 liter. Add 1 g of CuSO^ and 0.5 mL
of H2S0^ to preserve.
1.2 Intermediate phenol solutions: Dilute appropriate volumes of the
stock phenol solution to obtain working phenol concentrations of
10 |Jg/roL and 1 pg/mL.
1.3- Buffer solution: Dissolve 16.9 g of NH^Cl in 143 mL of concen-
trated NH.OH and dilute to 250 mL with distilled water. Two mL of
buffer solution should adjust 100 mL of sample to pH 10.
1.4 Aminoantipyrine solution: Dissolve 2 g of 4-aminoantipyrine in
distilled water and dilute to 100 mL.
1.5 Potassium ferricyanide solution: Dissolve 8 g of potassium ferri-
cyanide in distilled water and dilute to 100 mL.
1.6 Chloroform
2. Procedure
2.1 Prepare a series of standards with phenol concentrations ranging
from 0.004 to 0.040 |Jg/mL using the intermediate phenol solutions
(1.2). Total working volume of each standard must be at least
500 mL.
2.2 To 500 mL of standards and samples in separatory funnels, add
10 mL of buffer solution (1.3) and mix. Adjust the pH to 10 + 0.2
by adding additional buffer if necessary.
2.3 Add 3.0 mL of 4-aminoantipyrine solution (1.4) and mix.
2.4 Add 3.0 mL of potassium ferricyanide solution (1.5) and mix.
2.5 After 3 minutes, extract with 25 mL of chloroform (1.6). Shake
the separatory funnel at least 10 times, let the CHCl^ settle,
shake again 10 times, and let the layers separate.
92.
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T'r'r'iWC G'JiD^ SHcf.i
2.6 Filter chloroform extracts through filter paper into 50-mL beakers
and cover with watchglasses.
2.7 Read the absorbance of the samples and standards against the blank
at 460 nm.
:3. .Calculations .
3.1 Prepare standard curve by plotting the absorbance value of
standards versus the corresponding phenol concentrations.
3.2 Obtain the concentration value of the sample directly from the
standard curve.
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DETERMINATION OF PHENOL: OXIDATIVE COUPLING REACTION OF
3-METHYL-2-BENZOTHIAZOLINONE HYDRAZONE WITH PHENOLS (9)
1. Reagents
1.1 Stock phenol solution: Dissolve 1 g of phenol in 500 mL of dis-
tilled water and dilute to 1 liter. Add 1 g of CuSO^ and 0.5 mL
of H^SO^ to preserve.
1.2 Intermediate phenol solutions: Dilute appropriate volumes of the
stock phenol solution to obtain working phenol concentrations of
10 (Jg/mL and 1 (Jg/mL.
1.3 3-methyl-2-benzothiazolinone hydrazone solution, 0.05%: Dissolve
0.1 g of 3-methyl-2-benzothiazolinone hydrazone hydrochloride in
200 mL of distilled water.
1.4 Ceric ammonium sulfate solution: Add 2.0 g of eerie ammonium
sulfate and 2.0 mL of concentrated sulfuric acid to 150 mL of
distilled water. After the solid has dissolved, dilute to 200 mL
with distilled water.
1.5 Buffer solution: Dissolve in the following order: 8.0 g of sodium
hydroxide, 2.0 g of EDTA (disodium salt), and 8.0 g of boric acid
in 200 mL of distilled water. Dilute to 250 mL with distilled
water.
1.6 Working buffer solution: Mix equal volumes of buffer solution
(1.5) and ethanol.
1.7 Chloroform
2. Procedure
2.1 Prepare a series of standards with phenol concentrations ranging
from 0.004 (Jg/mJ. to 0.060 pg/mL, using the intermediate phenol
solutions (1.2). Total working volume of each standard must be
at least 500 mL.
2.2 To 500 mL of standards and samples in separatory funnels, add 4 mL
of 3-methyl-2-benzothiazolinone hydrazone solution (1.3) and mix.
2.3 After 5 minutes, add 2.5 mL of ceric ammonium sulfate solution
(1.4) and mix.
2.4 After an additional 5 minutes, add 7 mL of working buffer solution
(1.6) and mix. 1
2.5 After 15 minutes, add 25 mL of chloroform. Shake the separatory
funnel at least 20 times. Allow the layers to separate, and pass
the chloroform layer through filter paper into 50-mL beakers and
cover with watchglasses.
.94.
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TVBtfiG GUtP-F. SHIltT
2.6 Read the absorbance at 490 tun against a reagent blank.
3. Calculations
3..1 Prepare standard curve by plotting absorbance against concen-
- tration values.
3.2 Obtain the sample concentration directly from the standard curve.
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TY?iMG GolDr fiiUff
DETERMINATION OF PHENOL: ULTRAVIOLET DETERMINATION OF PHENOL
UTILIZING THE SPECTRO PRODUCTS MODEL PH-3 PHENOL INSTRUMENT (146)
1. Reagents
1.1 Stock phenol solution: Dissolve 1 g of phenol in 500 mL of dis-
-tilled water and dilute to 1 liter. Add 1 g of CuSO^ and 0.5 mL
of H^SO^ to preserve.
1.2 Intermediate working phenol solutions: Dilute appropriate volumes
of the stock phenol solution to obtain working solutions contain-
ing 1.0 (Jg/mL and 0.5 (Jg/mL of phenol.
1.3 Sodium hydroxide solution, 4M: Dissolve 16 g of NaOH in 75 mL of
distilled water. Dilute to 100 mL after the solid has dissolved.
1.4 Phosphoric acid solution, 22%: Dilute 30 mL of 75% phosphoric
acid to 100 mL with distilled water.
2. Procedure
2.1 Set up and calibrate the spectrophotometer according to the oper-
ator's manual. Clean the inside of the 10 cm absorption cell by
rinsing several times with phenol-free distilled water.
2.2 Prepare a series of phenol standards in the range 0.002 to
0.10 (Jg/mL by diluting appropriate volumes of the intermediate
working solutions (1.2) with phenol-free water. Include a stan-
dard blank. Adjust the pH of the standards to 4.0 by adding
phosphoric acid solution (1.4), drop-by-drop.
2.3 Rinse the 10 cm cell with the standard blank twice, then fill the
cell with the blank solution. Place the cell inside the sample
compartment.
2.4 Adjust the coarse Lamp "A" control to a zero (0) reading on the
meter. Set the scale expand to the lOx position and adjust the
fine Lamp "A" control to obtain a zero (0) reading on the meter.
Return scale expand switch to the lx position. Open the sample
compartment and remove the sample cell. Handle the cell in such a
way as not to get fingerprints on it.
2.5 Add approximately 2 drops of sodium hydroxide solution (1.3) to
the cell. Replace the cell in the sample compartment, taking care
to be sure that the cell is returned to EXACTLY the original
position.
2.6 Observe the meter reading. Set the scale expand switch to lOx and
record the reading.
96
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TMNG GUIDE SHEET
2.7 Repeat the steps 2.3 to 2.6 for the phenol standards, beginning
with the lowest concentration-and working up to the higher con-
centrations.
3. Calculations
3.1. Divide the meter readings obtained for the phenol standards and
samples by 10.
3.2 Plot the corrected meter readings (2.1) on semi-logarithmic graph
paper against the known concentrations of the phenol standards.
3.3 Obtain the concentration value of the samples directly from the
standard curve, using the corrected meter readings (3.1).
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mMMCi G'.I.DE SHElir
DETERMINATION OF PHENOLS: ULTRAVIOLET DETERMINATION OF TOTAL PHENOLS (119)
1. Reagents
1.1 Stock £-cresol solution: Dilute 1.0 g of £-cresol to 100 mL with
distilled water. 1 mL = 10 mg £-cresol.
1.2 Intermediate £-cresol solutions: Dilute appropriate volumes of the
stock £-cresol solution (1.2) to obtain intermediate working
£-cresol solutions containing 1 !0 |Jg/mL and 10 pg/mL p-cresol.
1.3 Sodium hydroxide solution, AM: Dissolve 16.0 g of sodium hydrox-
ide in 500 mL of distilled water. Dilute to 1 liter.
1.4 Phosphoric acid solution, 22%: Dilute 30 ml of 75% phosphoric
acid to 100 mL with distilled water.
2. Procedure
2.1 Prepare a series of £-cresol standards in the range 1 to 30 pg/rnL
using the intermediate £-cresol solutions (1.2).
2.2 Take a measured portion of the standards and samples and adjust
the pH to 4 with phosphoric acid solution. Read the absorbance
of the standards and samples at 293 nm on an ultraviolet spectro-
photometer.
2.3 Remove the cell from spectrophotometer, adjust the pH of the
solution to 12 by adding a few drops of sodium hydroxide solution
(1.3). Place the cell in the spectrophotometer and read the
absorbance.
3. Calculations
3.1 Subtract the absorbance of samples and standards at pH 4 from the
absorbance at pH 12. Prepare a calibration curve by plotting the
difference in absorbance versus concentration values of the
£-cresol standards.
3.2 The concentration of total phenolic compounds in the sample is
obtained directly from the calibration curve.
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Xi?\h£ GL'iDn SiirPI
APPENDIX D
METHOD FOR ANALYSIS OF SOLID AND SEMISOLID
ENVIRONMENTAL SAMPLES FOR TOTAL PHENOLIC COMPOUNDS
PHENOLICS, TOTAL RECOVERABLE
1. Scope and Application
1.1 This method is applicable to the analysis of solid and semisolid
environmental samples.
1.2 The method is capable of measuring total phenolic compounds at
concentrations greater than 50 ng/g using the direct colorimetric
method or at the 5 ng/g level by the chloroform extraction pro-
cedure.
2. Summary of Method
2.1 Phenolic compounds are extracted from solid and semisolid samples
by blending with dilute sulfuric acid solution. Dichloromethane
is used to extract the phenols from the acid solution. The phe-
nolic compounds are extracted from dichloromethane into dilute
sodium hydroxide solution. The phenolic compounds present in the
sodium hydroxide solution are quantified by the 4-aminoantipyrine
colorimetric procedu^s.
3. Comments
3.1 The extraction efficiency and absorption maxima are different for
various substituted phenolic compounds. Because it is not possi-
ble to duplicate the mixture of phenolic compounds expected in a
variety of environmental samples, phenol has been selected as the
standard compound for quantification.
4. Sample Handling and Preservation
4.1 Samples should be collected in wide mouth glass bottles which have
been cleaned in a solution of chromic acid.
4.2 Samples should be frozen as sojn as possible after collection.
Frozen samples may be stored for up to 2 weeks without significant
loss of phenol.
5. Interferences
5.1 Compounds which absorb at the analytical wavelength may produce a
positive interference.
.99.
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IV^AJO GU!DE SHi'i'T
6'. Apparatus
6.1 Blender suitable for laboratory applications
6.2 Beakers, 250 mL
-6.3- Plastic centrifuge bottles, 250 mL
6.4 Centrifuge
6.5 Separatory funnels, 250 ml and 500 mL
6.6 Flasks, volumetric, 100 mL
6.7 Conical filter funnels
6.8 Glass fiber filter paper
6.9 _ Spectrophotometer, for use at 460 nm or 510 nm
6.10 pH meter or pH paper with pH range 3.0 to 5.5 and pH range
9.0 to 12.0
7. Reagents
7.1 Sodium sulfate, anhydrous powder
7.2 Sulfuric acid solution, 0.18M: dilute 10 mL of concentrated
H^SO^ to 1 liter with distilled water
7.3 Dichloromethane
7.4 Sodium hydroxide solution, 0.005N: Dissolve 200 mg of NaOH in
distilled water and dilute to 1 liter.
7.5 Stock phenol solution: Dissolve 1.0 g of phenol in freshly
boiled and cooled distilled water and dilute to 1 liter.
1 mL = 1 rag phenol.
7.6 Working solution A: Dilute 10 mL of stock phenol solution to
1 liter with distilled water. 1 mL = 10 mg phenol.
7.7 Working solution B: Dilute 10 mL of working solution A to 1 liter
with distilled water. 1 mL = 0.1 mg phenol.
7.8 Phosphoric acid solution, 1 + 9: Dilute 5 mL of 85% phosphoric
acid to 50 mL with distilled water.
7.9 Buffer solution: Dissolve 16.9 g of NH^Cl in 143 mL of concentra-
ted NH^OH and dilute to 250 mL with distilled water. Two mL
should adjust 100 mL of sample extract to pH 10.
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7.10 Aminoantipyrine solution A: Dissolve 2 g of 4-aminoantipyrine- in
distilled water and dilute to 100 ml.
7.11 Aminoantipyrine solution B: Dissolve 0.4 g of 4-aminoantipyrine
in distilled water and dilute to 100 mL.
7.12 Potassium ferricyanide solution A: Dissolve 8 g of K^FeCCN)^ in
distilled water and dilute to 100 mL.
7.13 Potassium ferricyanide solution B: Dissolve 1.6 g of K Fe(CN)^ in
distilled water and dilute to 100 mL.
7.14 Chloroform
Procedure
8.1 Acid Extraction
,8.1.1 Place 25 to 50 g of frozen sample in the blender.
Blend quickly to a powder.
8.1.2 Weigh a portion of sodium sulfate (7.1) equivalent to-
the weight of the sample amd add to the sample in the
blender and mix.
8.1.3 Add 100 mL of sulfuric acid solution (7.2) to the dry
mixture. Blend at high speed for 5 minutes.
8.1.4 Transfer the contents of the blender into a plastic
centrifuge bottle. Rinse the blender with two 10-mL
portions of the sulfuric acid solution (7.2) and
transfer the washings to the centrifuge bottle.
8.1.5 Centrifuge the mixture at moderate speed for 10 minutes.
Decant the supernatant fluid from the centrifuge bottle
to a 500 mL separatory funnel. Do not disturb the
residue at the bottom of the centrifuge bottle.
8.2 To the supernatant acid extract in the 500 mL separatory funnel,
add 50 mL dichloromethane (7.3) and extract by shaking for 1 minute.
Allow the layers to separate and transfer the dichloromethane to a
250-mL separatory funnel. If the dichloromethane extract is
turbid, filter through glass fiber filter paper into the
250 mL separatory funnel.
8.3 Extract the aqueous layer two more times using two 25-mL
volumes of dichloromethane, shaking for 1 minute, and separating
the layers. Combine all the dichloromethane extracts in the
250-mL separatory funnel, filtering if necessary.
101
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GUtDc SH^T
8.4 If an emulsion forms during'the extraction with dichloromethane,
combine the emulsified layers in a centrifuge bottle. Centrifuge
at moderate speed for 5 minutes'. Separate the layers and transfer
the dichloromethane to the separatory funnel, filtering through
glass fiber filter paper if necessary.
>8.5 Add 30 raL of sodium hydroxide solution (7.4) to the dichloro-
methane in the separatory funnel. Extract the paenolic compounds
from the dichloromethane by shaking for 1 minute. Filter the
sodium hydroxide solution through glass fiber filter paper into a
100-mL volumetric flask.
8.6 Extract the dichloromethane two more times using 15 mL of the
sodium hydroxide solution (7.4) for each extraction. Filter and
combine the sodium hydroxide extracts in the volumetric flask.
Dilute to 100 mL with sodium hydroxide solution (7.4).
8.7 If an emulsion forms during the sodium hydroxide extraction,
separate the emulsified layers and combine these in a centrifuge
bottle. Centrifuge at moderate speed for 5 minutes, separate the
layers, and filter the sodium hydroxide solution through glass
fiber filter paper into the volumetric flask. Dilute to 100 mL
with sodium hydroxide solution (7.4).
8.8 Direct photometric method
8.8.1 Using working solution A (7.6) prepare the following
standards in 100 mL volumetric flasks. Use sodium
hydroxide solution (7.4) to dilute to volume.
mL working solution A concentration (pg/1)
0 0
0.5 50
1.0 100
2.0 200
5.0 500
7.0 700
10.0 1000
20.0 2000
8.8.2 Transfer 100 mL of the sodium hydroxide sample extract
or an aliquot diluted to 100 mL with sodium hydroxide
solution (7.4) into a 250 mL beaker. Transfer the total
volume of standards to beakers.
8.8.3 Adjust the pH of samples and standards to 4.by. adding
phosphoric acid solution (7.8) drop-by-Irop.
8.8.4 Add 2.0 mL of buffer solution (7.9) and mix. The pH of
the sample and standards should be 10 + 0.2.
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8.8.5 Add 2 mL of aminoantipyrine solution A (7.10) and mix.
8.8.6 Add 2 mL of potassium ferricyanide solution A (7.13) and
mix.
8.8.7 After 15 minutes, read absorbance at 510 nm.
8.9 Chloroform extraction method
8.9.1 Using working solution B (7.7) prepare the
following standards in 100 mL volumetric
flasks. Dilute to volume with sodium
hydroxide solution (7.4).
mL of working solution B concentration (ng/L)
0 0
5 5.0
10 10.0
20 20.0
30 30.0
40 40.0
8.9.2 Transfer the standards and 100 mL of the sodium
hydroxide sample extract into 250-mL separatory funnels.
8.9.3 Adjust, the pH to 4 by adding phosphoric acid solution
(7.8) drop-by-drop.
8.9.4 Add 2.0 mL of buffer solution (7.9) and mix. The pH of the
sample and standards should be 10 ± 0.2.
8.9.5 Add 3.0 mL of aminoantipyrine solution B (7.11) and mix.
8.9.6 Add 3.0 mL of potassium ferricyanide solution B (7.13) and
mix.
8.9.7 After 3 minutes, extract with 15 mL of chloroform (7.14).
Shake the separatory funnel at least 10 times, let the
chloroform settle, shake again 10 times, and let chloroform
settle again.
8.9.8 Filter chloroform extracts through glass fiber filter paper.
Do not add more chloroform.
8.9.9 Read the absorbance of the samples and standards against a
chloroform blank at 460 nm.
Calculation
9.1 Prepare a standard curve by plotting the absorbance value of
standards versus the corresponding phenol concentrations.
103
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TvPIPiv- GUI Oil SHEF.V
or.'.;;,:;-,
CF- PACr;
9.2 Obtain the conceatration value of sample directly from the
standard curve.
9.3 Calculate the concentration of total phenolic compounds in the
solid sample using thi. following equation:
"Mg/8 = A x 0.1
B
where A = concentration of sample as read from calibration curve
(pg/L)
B = weight of solid sample used for extraction (g)
:104
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