DOT/FAA/AM-00/16

Nicole T. Vu

Arvind K. Chaturvedi

Dennis V. Canfield

John W. Soper

Civil Aeromedical Institute

Federal Aviation Administration

Oklahoma City, Oklahoma 73125

Doris M. Kupfer

Bruce A. Roe

Department of Chemistry and Biochemistry

University of Oklahoma

Norman, Oklahoma 73019

May 2000

Final Report

This document is available to the public

through the National Technical Information

Service, Springfield, Virginia 22161.

DNA-Based Detection of EthanolProducing Microorganisms in

Postmortem Blood and Tissues by

Polymerase Chain Reaction

Office of Aviation Medicine

Washington, D.C. 20591

U.S. Department

of Transportation

Federal Aviation

Administration
NOTICE

This document is disseminated under the sponsorship of

the U.S. Department of Transportation in the interest of

information exchange. The United States Government

assumes no liability for the contents thereof.
i

Technical Report Documentation Page

1. Report No. 2. Government Accession No. 3. Recipient's Catalog No.

DOT/FAA/AM-00/16

4. Title and Subtitle 5. Report Date

DNA-Based Detection of Ethanol-Producing Microorganisms in Postmortem May 2000

Blood and Tissues by Polymerase Chain Reaction 6. Performing Organization

Code

7. Author(s) 8. Performing Organization

Report No.

Vu, N.T., Chaturvedi, A.K., Canfield, D.V., Soper, J.W.1,

Kupfer, D.M., and Roe, B.A.2

9. Performing Organization Name and Address 10. Work Unit No. (TRAIS)

1

FAA Civil Aeromedical Institute

P.O. Box 25082

Oklahoma City, Oklahoma 73125

2

Department of Chemistry and Biochemistry

University of Oklahoma

Norman, Oklahoma 73019

12. Sponsoring Agency name and Address 13. Type of Report and Period

Covered

Office of Aviation Medicine

Federal Aviation Administration

800 Independence Ave., S.W.

Washington, D.C. 20591 14. Sponsoring Agency Code

15. Supplemental Notes

This work was accomplished under the approved tasks AM-B-97-TOX-202, AM-B-98-TOX-202, AM-B-99-

TOX-202, and AM-B-00-TOX-202.

16. Abstract

Forensic investigation of fatal aircraft accidents usually includes the analysis of biological samples for ethanol to

establish if alcohol intoxication is a factor in the accidents. The quantitative aspects of ethanol are often

complicated by postmortem putrefactive changes, leading to microbial fermentation-mediated production of

alcohol and its subsequent redistribution. Without establishing the ethanol origin (antemortem consumption or

postmortem production), a precise interpretation of the alcohol analytical results remains a challenge.

Therefore, a DNA-based assay was developed using the polymerase chain reaction and microbial DNA primers

designed for identifying 3 commonly encountered ethanol-producing microorganismsCandida albicans,

Proteus vulgaris, and Escherichia coli. The present study focused on examining the applicability of the microbial

DNA primers in establishing the existence of postmortem alcohol in samples. The results suggested that speciesspecific primers could be employed to identify ethanol-producing microorganisms in forensic samples without

requiring bacterial cultivation. Continued studies are warranted to define additional primer sequences that are

distinctive for ethanol-producing microorganisms.

17. Key Words 18. Distribution Statement

Polymerase Chain Reaction, PCR, DNA Analysis, DNA

Primers, Oligonucleotides, Postmortem Alcohol, Ethanol,

Putrefaction, Microorganisms, Fermentative

Microorganisms, Candida Albicans, Proteus Vulgaris,

Escherichia Coli

Document is available to the public through the

National Technical Information Service

Springfield, Virginia 22161

Security Classif. (of this report) 20. Security Classif. (of this page) 21. No. of Pages 22. Price

Unclassified Unclassified 13

Form DOT F 1700.7 (8-72) Reproduction of completed page authorized

1

DNA-BASED DETECTION OF ETHANOL-PRODUCING MICROORGANISMS IN

POSTMORTEM BLOOD AND TISSUES BY POLYMERASE CHAIN REACTION

INTRODUCTION

Although use of alcoholic beverages in moderation

is generally acceptable for recreational purposes, it is

irrefutable that alcohol abuse is a social and medical

problem of global significance. It is estimated that

two-thirds of all adults in the United States are

occasional drinkers; about 12% among them are

heavy users with risk of becoming alcohol dependent

(1). The primary acute effect of ethanol is central

nervous system depression, which leads to increased

reaction time, diminished fine motor control, and

impaired critical faculty (2). The direct hazards of

such effect are manifested in the statistics of fatal

motor vehicle accidents, with ethanol as a major

contributing factor in 50% of all cases (3). The legal

limit of blood ethanol for motor vehicle operators in

most states is 0.08 or 0.10% (w/v), but it is considerably lower for pilots. According to the Federal

Aviation Regulation (FAR) 91.17, no person may

operate or attempt to operate an aircraft with a blood

ethanol content of 0.04% or greater (4). However,

ethanol concentrations exceeding the FAR limit were

found in at least 6% of fatal aircraft accident cases

during a 2-year (1989-1990) period (5).

Forensic investigation of fatal aircraft accidents usually includes the determination of ethanol concentration in biological samples collected during autopsy

from pilots to establish if alcohol intoxication is the

cause of the accidents. However, postmortem putrefactive changes—microbial fermentation-mediated

production of alcohol and its subsequent redistribution—often complicate the interpretation of determined

ethanol levels. Generally, between 12 to 57% of ethanol-positive cases were attributed to postmortem

production (3,5). There were other instances in which

a significant amount of ethanol was found, but the

origin (antemortem ingestion or postmortem production) of the alcohol was undetermined. This

situation is emphasized in a study of aviation accidents that occurred during 1989-1990, where no

determination could be made regarding the origin of

ethanol in 45% of all positive cases (5). The lack of

direct evidence for antemortem ingestion, coupled

with putrefaction effects, requires interpretation of

postmortem ethanol concentrations based on the

atypical distribution of this substance in different

biological compartments. This normally requires

analysis of at least 2 types of specimens, such as blood

(or tissues) and vitreous humor, urine, or bile (3,5,6).

However, these specimens are often not available.

Recently, an altered urinary concentration ratio of 5-

hydroxytryptophol (5-HTOL) to 5-hydroxyindole3-acetic acid (5-HIAA) has been suggested as a possible

indicator of ethanol consumption (7,8). At present,

this test can be performed on only urine specimens,

as the reliability of this ratio has not yet been established in other biological fluids.

In a recent Federal Aviation Administration (FAA)-

supported study, Kupfer et al. (9) described the

development of species-specific DNA primers for

selected microorganisms with ethanol-producing capability. The premise was that the presence of the

ethanol producers in putrefying tissues constitutes a

possible indication of the microbial origin of the

alcohol in the samples. While viability of the microbial species is crucial to the culturing techniques,

DNA-based methods allow the identification of the

microbial contaminants, whether or not they survive

the conditions of environment and/or the specimens’

preservation and storage. Here, the polymerase chain

reaction (PCR) assay was easily performed, and the

results were quantifiable, using oligonucleotide primers for 3 commonly encountered ethanol-producing

microorganisms—Escherichia coli, Proteus vulgaris,

and Candida albicans. Detailed information on the

development of the primer pairs, along with the

protocols for the genomic DNA isolation and amplification, has been previously reported (9).

The objective of the present study was to examine

the applicability of the aforementioned microbial

DNA primers in establishing the existence of postmortem ethanol. The question was whether a PCRbased assay using species-specific primers could
2

effectively determine the origin of ethanol present in

certain forensic samples. Therefore, 3 batches of

specimens with indisputable ethanol interpretation,

based on both typical and atypical distribution criteria, were selected for the PCR analysis.

MATERIALS AND METHODS

Materials

For alcohol analysis, all reagents were of analytical

grade and solvents were of chromatographic grade.

The reagents and solvents employed in the DNA

analysis were of molecular biology grade or sterilized

by autoclaving. These reagents, solvents, standards,

internal standards, and other necessary supplies were

obtained from commercial sources. The QIAamp®

Tissue Kit for DNA isolation was obtained from

QIAGEN, Inc., Valencia, CA.

Description of Postmortem Forensic Samples

The forensic specimens were collected during autopsy and placed in the FAA’s TOX-BOX evidence

containers. The boxes containing samples were

shipped with ice packs for analysis to the FAA’s Civil

Aeromedical Institute (Oklahoma City, OK) by an

air carrier service for next-day delivery. Blood was

submitted in 10-mL vacutainers, containing 10 mg

of sodium fluoride and 20 mg of potassium oxalate.

Tissue specimens were submitted in sterile plastic

bags with no added preservatives. Using standard

laboratory procedures, these samples were analyzed

for drugs, alcohols, carboxyhemoglobin, and cyanide, and stored at –20o

C. Alcohols were identified

and quantified using headspace gas chromatography

with t-butanol as the internal standard. All cases with

a blood ethanol concentration equal to or greater

than 0.04% (40 mg/dL) were considered positive.

Other volatiles were screened at a limit of detection

of 1 mg/dL.

The criteria for determining postmortem alcohol

production were based on the absence of ethanol in

the vitreous humor and/or urine and positive ethanol

in the blood or tissues. In accordance with the results

of the alcohol analysis, the specimens were divided

into 3 batches. Batch 1 consisted of 23 blood and

tissue samples wherein no ethanol was detected, Batch

2 was composed of 24 blood and tissue samples where

the presence of ethanol was attributed to consumption, and Batch 3 was composed of 22 samples

containing ethanol due to postmortem production.

Solid tissues or tissue fluids, including brain, heart,

kidney, liver, lung, and muscle, were used, depending on the specimen availability.

Microbial Cell Culture and Concentration

Determination

Isolates of Escherichia coli (strain ATCC 8739),

Proteus vulgaris (University of Oklahoma strain

10052), and Candida albicans (strain ATCC 10231)

were used for the preparation of stock culture for each

species. The microbial cells were obtained from overnight broth culture and maintained at –70°C in the

presence of 17% glycerol. To estimate the concentration of the microbes in a sample, a calibration curve

was constructed for each species to correlate the

colony-forming units (cfu) and the absorbance at 600

nm of the cell suspension in 10 mM MgSO4

. The

bacteria, E. coli and P. vulgaris, were grown in Lbroth (1% peptone, 0.5% yeast extract, and 0.5%

NaCl in water), while C. albicans was cultured in

modified Saubouraud’s medium (1% neopeptone

and 2% dextrose in water). Each 10-mL broth culture

was incubated at 37°C with aeration to late log phase.

The microbial cells were collected by centrifugation

and resuspended in 10 mL of sterile 10 mM MgSO4

solution. The cell suspensions were sonicated (300

Ultrasonik, NEY Barkmeyer Division, Yucaipa, CA)

at 50% power for 60 sec to separate the cell clumps.

A serial dilution was prepared for each cell suspension; a 100-μL sample was removed from each series,

and it was streaked on the appropriate agar plate for

a colony plate count following 24-hour incubation at

37°C. The E. coli and P. vulgaris cells were spread on

L-broth agar and MacConkey’s agar, respectively. C.

albicans cells were spread on Saubouraud’s agar plates.

Absorbance of each diluted sample was measured at

600 nm (Vectra XM, Hewlett Packard, Waldbronn,

Germany). Regression analysis of the respective absorbance vs. cfu/mL generated a representative calibration curve for the particular organism.

DNA Extraction

DNA was extracted according to the protocol

described in the previous study (9). Briefly, cells from

300 μL of the blood or tissue fluid samples were

collected by centrifugation at 8,000xg for 3 min in a

microcentrifuge (235C, Fisher Scientific Co., Pittsburgh, PA). For solid tissues, approximately 0.03 cm3

(≈ 30 mg) of each sample was grated, using a sterile

scalpel and aseptic techniques, and then suspended in
3

270 μL of 10 mM MgSO4. Pellets were suspended in

Erythrocyte Lysis Solution (QIAGEN Inc., Valencia,

CA) followed by centrifugation, and then reconstituted in Cell Suspension Solution (Gentra Inc., Minneapolis, MN). To facilitate the lysis of the microbial

cell wall, the cell suspension was incubated at 37°C

for 70 min in the presence of 200 U lyticase (Sigma,

St. Louis, MO) and 0.5% β-mercaptoethanol (Fisher

Scientific Co., Pittsburgh, PA). DNA was isolated

using the QIAmp® Tissue Kit, as described in the

manufacturer’s protocol. Along with each batch of

samples, a negative blood control and a reagent blank

were processed, as well as 3 positive control samples

of E. coli, P. vulgaris, and C. albicans.

The positive control samples were prepared for

each microorganism by inoculating 200 μL of negative human blood (Oklahoma Blood Institute, Oklahoma City, OK) with 100 μL of the microbial cell

suspensions collected from overnight cultures. The

positive control samples contained 105

 cfu of the

bacteria or 103

 cfu of the yeast. Uninoculated human

blood and 10 mM MgSO4

 were used for negative and

reagent control in 300-μL aliquots, respectively.

DNA Amplification

To detect the target DNA in the extracts, PCR was

performed according to the protocol developed in the

earlier study (9). Previously designed oligonucleotide primers (9) were synthesized on a Beckman

Oligo 1000M DNA Synthesizer (Beckman Instruments, Houston, TX). Approximately 20 ng of the

extracted DNA (8 μL of the extract) from each sample

was added to 3 separate reaction mixtures that contained the primer pair specific for either C. albicans,

P. vulgaris, or other enteric bacteria. Those primers

were FAA37, 11 for C. albicans; FAA46, 45 for P.

vulgaris; and FAA30, 31 for E. coli, P. vulgaris, and

other enteric bacteria (9). The PCR was performed in

a Perkin-Elmer 9600 thermocycler (Perkin Elmer

Corporation, Norwalk, CT) in 35 cycles, following a

5-min denaturation at 95°C. Each thermal cycle

consisted of 3 steps: denaturation (94°C, 1 min),

primer annealing (50°C, 2 min), and primer extension (72°C, 3 min). The last cycle was followed by the

extension step at 72°C for 10 min and then cooling

to 15°C for storage. To inactivate the polymerase, 5

μL of 200 mM EDTA was added to each of the

amplified samples and stored at –20°C until used.

Semiquantitation of Microbial Cells by Duplex

PCR

Optimization for the duplex amplification was

conducted using E. coli as the representative organism and C. albicans as the internal control. The

control template was prepared from a pool of the

extracts obtained with a blood matrix containing

3.0x105

 cfu C. albicans and then diluted in 2 parts of

sterile water. The diluted extract of the control template (8 μL; ≈20 ng DNA) was added to each reaction

mixture. The E. coli template was prepared from a

series of blank blood that had been supplemented

with 105

, 106, and 107 microbial cells. The PCR

conditions were optimized with respect to the amounts

of the Taq-polymerase (M186B, Promega, Madison,

WI), the MgCl2

 buffer (M188J, Promega, Madison,

WI), and the dNTPs (100 mM, Pharmacia Biotech,

Piscataway, NJ). The thermal cycles and the primer

concentration for each template were the same as

previously described in the single target sequence

amplification.

Estimation of the microbial concentration in the

forensic samples was accomplished by performing

the duplex PCR for selected samples under optimized

conditions. The E. coli calibration curve was examined in the range from 103

 to 5.5x107 cfu/300 μL.

The assay precision was investigated in 5 replicates,

using the same extract from 6 E. coli standards,

ranging from 103

 to 5.5x107 cfu/300 μL, which were

analyzed on separate occasions. Precision was evaluated by means of the normalized ratios obtained from

the extrapolated cfu values for E. coli and C. albicans.

PCR Product Gel

Gel electrophoresis was performed to identify the

PCR products using a Horizontal Mini-Gel Electrophoresis Unit (Sigma-Aldrich, St. Louis, MO). The

10x7.5x0.4-cm gels were cast from a solution of 2%

agarose in 0.5X TBE buffer (45 mM Tris, 45 mM

borate, and 1 mM EDTA) and 0.5 μg/μL ethidium

bromide (Sigma, St. Louis, MO). Each 8 μL of the

amplified sample was mixed with 2 μL of the gel

loading buffer (20 mM Tris, 2.5 mM EDTA, 50%

glycerol, and 0.2% bromophenol blue). This mixture

was electrophoresed in parallel with a 123 Base Pair

DNA ladder (Life Technologies Inc., Gaithersburg,

MD). Electrophoresis was carried out at 105 volts

(DPS 1000, Buchler Instrument Inc., Fort Lee, NJ)
4

for approximately 1.5 hours, at which time the bromophenol blue dye front had traveled at least 7.5 cm

from the wells. Subsequently, the gels were photographed under UV-illumination (U.V.P. Inc., San

Gabriel, CA), using a red 23A filter (Tiffen Manufacturing Corp., Hauppage, NY). The intensity of the

DNA bands represents the relative amount of specific

amplification product, when compared with standard samples of known concentration.

RESULTS

A typical calibration curve for determining the

concentration of the microbial cell suspensions in the

MgSO4

 solution is presented in Fig. 1. The calibration curves were linear over the range examined with

r2

 = 0.986 for E. coli, 0.997 for P. vulgaris, and 0.960

for C. albicans. The colony count approximated the

number of viable bacteria or yeast cells, while the

absorbance represented the amount of viable and

unviable microbial cells.

The PCR-products obtained from the positive

control samples of the 3 microorganisms are shown

in triplicate (Fig. 2). C. albicans yielded a 704 bp

fragment at 103

 cfu. The 384 bp and 1226 bp

Fig. 1. The calibration curve for E. coli

suspended in 10 mM MgSO4. A typical

linear curve (r2 = 0.986) used for determining the concentration of viable cells/

mL of the microorganism is shown. Absorbance was measured at 600 nm. cfu:

colony-forming units.

fragments were produced with E. coli and P. vulgaris

at 105

 cfu, respectively. The detectable levels for each

species in a mixed microbial blood matrix were determined to be 103

 cfu/300 μL (not shown for E. coli and

P. vulgaris). These levels were detected in 70% of the

tested control samples.

The forensic samples from all 3 batches were

screened using the 3 DNA primer pairs in separate

amplification reactions. Preliminary screening results indicated that amplicons produced by the primer

pair FAA30, 31 were detectable with high intensity in

samples of Batch 1 and Batch 3, while only 13% of

samples in Batch 2 (ethanol due to ingestion) were

positive (Table 1). C. albicans was detected in 9 to

33% of the samples from all the batches. P. vulgaris

was found only in Batch 3 samples, at a frequency of

less than 10%.

M 2 3 4 M 6 7 8 M 10 11 12

Fig. 2. The PCR products obtained from

representative microorganisms using

specific primer pairs. Lanes 2, 6, 10 C.

albicans (704 bp); lanes 3, 7, 11 E. coli

(384 bp); lanes 4, 8, 12 P. vulgaris (1226

bp). Marker lanes (M).
5

Since a significant number of samples with (Batch

3) and without (Batch 1) ethanol showed positive

results with the primer pair FAA30, 31 (Table 1), it

was suggested that the relative amounts of the microorganisms were responsible for the observed differences in the ethanol content between the 2 batches. A

semi-quantitative assessment of the microbial DNA

content among the samples was achieved by the

duplex PCR procedure. The goal was to determine

the relative numbers of the microorganisms present

in the forensic samples, both viable and unviable,

without reference to the exact number of their target

DNA templates. C. albicans template equivalent to

105

 cells was employed to normalize the efficiency of

the amplification process. In optimizing the duplex

PCR, the variables that consistently produced the

largest yield of the duplex targets, while remaining

within the linear amplification range, were selected

for use. Since the melting temperatures for primer

pairs FAA30, 31 and FAA37, 11 are similar (Tm ≈ 60-

64°C), their optimal annealing temperatures are also

equivalent (10). Hence, the conditions of the thermal

cycles and the concentration of the primers used for

the previous reaction (single template amplification)

were adopted for duplex priming. The concentrations of the polymerase, the dNTPs, and the MgCl2

buffer were the variables reexamined for assay optimization. The results confirmed the robustness of the

existing PCR condition. Accordingly, the polymerase

at 5 units, the dNTPs at 0.2 μmole, and MgCl2

 at 1.5

μmoles in a 100-μL final volume of the reaction

mixture were optimal for the duplex amplification. A

standard curve representing the range 103

-5.5x107

cfu/300 μL of E. coli was prepared by plotting the

ratio of the equivalent cfu for E. coli:C. albicans vs. E.

coli cfu (Fig. 3). The estimation of C. albicans cfu was

based on calibration samples prepared by amplifying,

in the absence of the E. coli template, the C. albicans

standards at the concentrations from 103

 to 105 cfu.

In examining Table 2 and Figs. 3 and 4, it is observable that the C. albicans and E. coli products accumulated with similar efficiencies, and the product ratios

in Table 2 and Fig. 3 demonstrate the linear quantitative range for the E. coli target (r2

 = 0.998). The

precision of the assay was assessed based on a relative

standard deviation (RSD) ≤ 20%, which indicated

the tolerable reproducibility of the quantitative estimation. A relative change in the PCR products could

be detected with a minimum of a 5-fold change of the

starting E. coli DNA template concentration.

A relative estimate of the microbial content in the

forensic samples was examined by using the primer

pair FAA30, 31. The bacterial concentrations in

samples from Batch 1 and Batch 3 were extrapolated

from the respective standard curve for E. coli obtained by the duplex PCR procedure (Fig. 3); the

results are given in Tables 3 and 4. Since the internal

control template in the duplex procedure was C.

Table 1. Percentage of samples in a particular batch where the targeted microbial

species were detected using the specific DNA primer pairs.

Percentage of samples*

Sample FAA37, 11 FAA46, 45 FAA46, 45

(for C. albicans) (for C. albicans) (for E. coli and other microbes)

Batch 1 samples Below detectable level 97%

(No ethanol detected)

26%

Batch 2 samples Below detectable level 13%

(Antemortem ethanol ingestion)

33%

Batch 3 samples 9% 9% 95%

(Postmortem ethanol production)

*The results are expressed in percentage of samples in which microbes under a particular category were

detected. The detectable levels for representative microbial species were 103

cfu.
6

Fig. 4. The amplicons obtained by duplex

PCR procedure from the standard

samples (also refer to Fig. 3). The top

amplicons are C. albicans, while the

lower amplicons are E. coli. C. albicans

template equivalent to 105 cfu was added

to the E. coli standards as the control

template. E. coli amplicons in lanes 1-7

represent 5.5x107, 107, 106, 105, 104, 103,

and 102 cfu/300 μL, respectively.

Fig. 3. Calibration curve for the estimation

of microbial content in selected samples

from Batch 1 and Batch 3 by a duplex PCR

procedure (r2 = 0.998). This curve was

prepared using E. coli standards as the

representative organism in the range from

103 to 5.5x107 cfu/300 μL; the internal

control template was prepared from C.

albicans at a constant concentration of 105

cfu. The cfu for C. albicans was obtained

by comparing the relative intensity of the C.

albicans product bands with that of the C.

albicans standard samples amplified in the

absence of E. coli. Each cfu ratio was

calculated based on the E. coli concentration in the standard sample and the respective estimated cfu for C. albicans product

(cfu: colony-forming units).

1 2 34 5 6 7
7

albicans, any forensic sample contaminated with C.

albicans was excluded from the assay. The target

DNA detectable by the FAA30, 31 primer set was

found at concentration levels correlating to 106

-108

cfu/mL in the majority of samples from both Batch 1

and Batch 3. The DNA concentration was 3 orders of

magnitude lower (≤ 103

 cfu) in about 10% of the

tested samples. The absence of both amplicons (target DNA and control DNA) in 5-17% of samples

suggests the reaction was inhibited, which might be

due to the putrefactive condition of those samples.

Thus, samples from Batch 1 and Batch 3 yielded

similar results when tested with the primer pair

FAA30, 31. These results indicate that ethanol production in Batch 3 was not related to any significant

difference in the amount of the microbial contamination (Mann-Whitney, p = 0.9399).

DISCUSSION

The specificity of the primers employed in this

study allowed the detection of 3 commonly encountered microbial speciesC. albicans, P. vulgaris, and

E. coli. As discussed previously (9), the FAA30, 31

primer pair was designed from a conserved region of

the 16S rDNA in the Enterobacteriaceae. Therefore,

the PCR products obtained by this primer pair could

provide an indication for the presence of any enteric

species, including Klebsiella, Yersinia, Samonella,

Proteus, and Escherichia. Many of these strains are

established ethanol producers (11).

The present results agree with the observation

made by Cory (11) that the greatest increases in

ethanol levels were usually associated not with yeast

contamination, but with enterococci or enteric bacilli.

*Ratio of the equivalent colony-forming unit for E. coli:C. albicans. The

equivalent cfu was derived from the evaluation of the relative intensity of the

PCR product bands. The E. coli product bands were compared with those of

the samples that made up the calibration curve (see Figs. 3 and 4). The C.

albicans product bands were compared with those of the C. albicans standard

samples amplified in the absence of E. coli. This same procedure was used in

evaluating the forensic samples. The mean values were obtained from 5

replicate analyses of the standard samples. The primer pairs used were FAA30,

31 and FAA37, 11. The C. albicans template equivalent to 105 cfu/mL was

added to each reaction as the internal control.

**RSD: relative standard deviation
8

Further, P. vulgaris was neither a major microbial

contaminant nor a significant source of postmortem

ethanol, based on the cases included in the present

study. The results also reveal that the primers FAA30,

31 have broad specificity, suggesting that these primers are not specific for the ethanol-producing enterobacteria. This observation is based on culturing

information obtained for selected samples of Batch 1,

which consisted of cases with no detectable ethanol.

While not all the microorganisms were identified, the

predominant species isolated were Streptococcus iniae,

Vibrio splendidus, Enterobacter aerogenes, and

Brochothrix thermosphacta. All of these species are

facultative anaerobes.Their major fermentative products are mixed acids, with ethanol as a minor component under appropriate condition (12,13). Thus, it is

likely that the DNA sequence flanked by the FAA30,

31 primer pair is conserved in one or more of the above

species and contributed the observed PCR product

bands signifying their presence. Perhaps the tissue conditions were unfavorable for ethanol production. Therefore, ethanol was not detected in these samples.

Whereas the broad specificity of FAA30, 31 allowed the detection of E. coli and many related

bacteria, the concentration of these species in most

samples from Batch 2 (ethanol levels due to antemortem consumption) was at or below the detectable

level of 103

 cfu/mL. The existence of putrefaction in

Batch 2 samples prompted the conclusion that species other than those characterized by the 3 primer pairs

were the predominant microorganisms, possibly including Staphylococci, Streptococci, Micrococcaceae,

Pseudomonas, Clostridium, Bacteroids, Peptococcus,

other yeasts, molds, etc. (11). The genomic DNA of
9

any of these predominant species would be extracted

by the protocol used but was not amplified during the

PCR. Therefore, designing additional PCR primers

specific for these other recognized ethanol producers

would expand the microbial detection range of this

technique.

SUMMARY

The 3 primers examined in this study could be

employed to identify microbial contaminants in putrid forensic samples. A positive result conferred by

the primer sets could lend credence to the microbial

origin of ethanol in postmortem alcohol production

cases. However, the use of microbial primers does not

distinguish other non-microbial sources of ethanol.

Additionally, the absence of PCR products resulting

from these primers does not exclude the presence of

other ethanol-producing microorganisms. The application of this DNA-based technique in resolving

the question of postmortem ethanol is pending the

development of additional primer sets specific for

other ethanol-producing bacteria and yeast.

Clearly, now that this approach has been shown to

be feasible, additional primers could be developed for

all implicated microbial species using unique ribosomal RNA gene regions as a source of primer sequences. Further studies should aim at expanding the

range of this DNA-based test for detecting the presence of ethanol-producing microorganisms.
10

REFERENCES

1. J.H. Jaffe, Drug addiction and drug abuse, in:

A.G. Gilman, T.W. Rall, A.S. Nies, P. Taylor

(Eds.), Goodman and Gilman’s the Pharmacological Basis of Therapeutics, 8th ed, Pergamon Press,

New York, 1990, pp. 538-9.

2. T.W. Rall, Hypnotics and sedatives; ethanol, in:

A.G. Gilman, T.W. Rall, A.S. Nies, P. Taylor

(Eds.), Goodman and Gilman’s the Pharmacological Basis of Therapeutics, 8th ed, Pergamon Press,

New York, 1990, pp. 370-9.

3. C.L. O’Neal, A. Poklis, Postmortem production

of ethanol and factors that influence interpretationa critical review, Am. J. Forensic. Med.

Pathol. 17 (1996) 8-20.

4. Alcohol or drugs, 14 C.F.R. Sect. 91.17 (1999).

5. D.V. Canfield, T. Kupiec, E. Huffine, Postmortem alcohol production in fatal aircraft accidents,

J. Forensic. Sci. 38 (1993) 914-7.

6. B.E. Stone, P.A. Rooney, A study using body

fluids to determine blood alcohol, J. Anal. Toxicol.

8 (1984) 95-6.

7. Helander, O. Beck, A.W. Jones, Distinguishing

ingested ethanol from microbial formation by

analysis of urinary 5-hydroxytryptophol and 5-

hydroxyindoleacetic acid, J. Forensic. Sci. 40 (1995)

95-8.

8. R.L. Hagan, A. Helander, Urinary 5-hydroxytryptophol following acute ethanol consumption: Clinical evaluation and potential

aviation applications, Aviat. Space. Environ.

Med. 68 (1997) 30-4.

9. D.M. Kupfer, A.K. Chaturvedi, D.V. Canfield,

B.A. Roe, PCR-based identification of postmortem microbial contaminants—a preliminary study,

J. Forensic. Sci. 44 (1999) 592-6.

10. J.S. Chamberlain, J.R. Chamberlain, Optimization of multiplex PCRs, in: K.B. Mullis, F. Ferré,

R.A. Gibbs (Eds.), The Polymerase Chain Reaction, Birkhäuser Boston, Cambridge, MA, 1994,

pp. 38-45.

11. J.E.L. Cory, Possible sources of ethanol ante- and

post-mortem: Its relationship to the biochemistry

and microbiology of decompositiona review, J.

Appl. Bacteriol. 44 (1978) 1-56.

12. Bergey’s Manual® of systematic bacteriology,

N.R. Krieg (Ed.), Willams & Wilkins Company,

Baltimore, MD, vol.1 (1984).

13. Bergey’s Manual® of systematic bacteriology,

P.H.A. Sneath (Ed.), Willams & Wilkins Company, Baltimore, MD, vol. 2 (1986).