APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Jan. 1986, p. 52-56 Vol. 51, No. 10099-2240/86/010052-05$02.00/0Copyright © 1986, American Society for Microbiology

Production of Toxin by Clostridium botulinum Type A StrainsCured of Plasmids

MICHAEL J. WEICKERT,' GLENN H. CHAMBLISS,l* AND HIROSHI SUGIYAMA2Department of Bacteriology1 and Food Research Institute,2 University of Wisconsin, Madison, Wisconsin 53706

Received 31 July 1985/Accepted 15 October 1985

Twelve strains of Clostridium botulinum type A and seven strains of Clostridium sporogenes were screened forplasmids by agarose gel electrophoresis of cleared lysates of cells from 5 ml of mid-log-phase culture. Nine typeA strains had one or more plasmids of 4.3, 6.8, or 36 megadaltons (MDa); several strains showed a largeplasmid of 61 MDa, but it was not consistently recovered. Four C. sporogenes strains had one or more plasmidsof 4.3, 5.6, or 36 MDa. Isolates obtained from cultures of plasmid-containing C. botulinum type A strainsgrown in ionic detergent broth and from spontaneously arising variants were screened both for toxinproduction and for plasmid content. Toxigenicity of C. botulinum could not be correlated with the presence ofany one plasmid.

The genetics of Clostridium botulinum toxin production isunknown except in the cases of toxin types C, and D, theproduction of which is mediated by specific bacteriophages(11, 12). The toxin type of these cultures is altered when aculture is cured to its resident tox+ bacteriophage andsubsequently reinfected with a phage specifying anothertoxin type (3). Reciprocal conversion of toxin production ofthis type also occurs when Clostridium novyi type A tox+bacteriophage infect phage-cured C. botulinum type C. Theresulting lysogenic strain is identical to C. novyi type Aincluding toxin production; however, it can be cured ofphage and converted to C. botulinum type C by reinfectionwith the type C tox+ phage (4).A prophage has been implicated in the production of type

A botulinum toxin (25), but such a relationship has certainlynot been firmly established. Since many C. botulinum type Astrains contain plasmids (20, 23; P. A. Kahn and M. D.Pierson, unpublished results), their toxigenicity might be dueto one or more of these plasmids. This is the case withClostridium tetani, in which a 50-megadalton (MDa) plasmidcontrols production of its neurotoxin (7), which is similar tobotulinum toxin in basic structure and biological activity(24).

In the present study, the plasmid contents of type Astrains were compared with those of Clostridium sporogenesstrains, an organism thought to be a nontoxigenic counter-part of the C. botulinum type A. The two species haveindistinguishable biochemical activities (17, 21), cross-reacting somatic antigens (19, 22), G-C ratios of 26 to 28%,and extensive RNA (13) and DNA (16, 18, 27) homology. Inaddition to comparison of the two clostridial strains, C.botulinum strains were cured of toxigenicity to see if theinduced effect correlated with plasmid loss.

MATERIALS AND METHODSOrganisms. C. botulinum type A strains were obtained from

the collection of the Food Research Institute, University ofWisconsin, Madison. The Af culture producing a mixture oftype A and F toxins (strain 84-SC2) was from D. Giminez,Mendoza, Argentina. C. sporogenes strains were from the

* Corresponding author.

Food and Drug Administration, Washington, D.C. Stockcultures were maintained in cooked meat medium (DifcoLaboratories, Detroit, Mich.) or brain heart infusion (BHI;Difco). Samples of exponentially growing cells that had beenquickly frozen in BHI broth containing 10% glycerol and thenstored at -80°C were used to inoculate fresh broth cultures.Clostridium strains were grown at 35°C in an anaerobic tent(model 12361; Coy Laboratory Products, Inc., Ann Arbor,Mich.) in an atmosphere of 80% N2, 10% H2, and 10% CO2.Media were prereduced for at least 8 h.

Escherichia coli V-517 and PRC122, used to obtain refer-ence plasmids, were from P. Vary, Northern Illinois Univer-sity, DeKalb. E. coli strains were grown aerobically at 37°Cin L broth containing, per liter, 10 g of tryptone (Difco), 10g of NaCl, and 5 g of yeast extract (Difco), and, whenneeded, 15 g of agar (Difco). Stocks were stored at -80°C inL broth containing 10% glycerol.Media and chemicals. Tryptone-peptone-glucose-yeast ex-

tract agar (5), glucose-yeast extract-peptone-thioglycolateagar (GYPT; 15), and gel diffusion agar (5) were prepared asdescribed elsewhere. Sodium deoxycholate (DOC) was ob-tained from Sigma Chemical Corp., St. Louis, Mo., andsodium dodecyl sulfate (SDS) was from Bio-Rad Laborato-ries, Richmond, Calif. Achromopeptidase (TBL-1) was fromWako Chemical, Dallas, Tex.; lysozyme (3 x crystallized,grade I) was from Sigma; Noble agar was from Difco;thiazine red was from Allied Chemical Corp., Morristown,N.J.; Merthiolate (Thimerosal) was from NutritionalBiochemicals Corp., Cleveland, Ohio; and RNase A (5 xcrystallized, type IA) was from Sigma.

Plasmid screening. Clostridial plasmids were extracted bya method based on that of Kahn and Pierson (unpublishedresults). A 0.2-ml volume of a culture incubated overnight inBHI medium was used to inoculate a second 5-ml BHIculture. After an 8-h incubation, the cells were harvested bycentrifugation at 12,000 x g for 15 min at 4°C. The pellet wassuspended in 50 Fal of Tris sucrose (50 mM Tris hydrochlo-ride, pH 8.0, with 25% [wt/vol] sucrose) and transferred to1.5-ml microfuge tubes (Walter Sarstedt, Inc., Princeton,N.J.). Two microliters of a lysozyme-TBL-1 solution (5 mgof each per ml of twice-distilled H2O) was added, and thesuspension was incubated at 37°C for 20 min.

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Cells were lysed by adding 380 ,ul of a freshly madealkaline-SDS solution (4% SDS, 20 mM EDTA, 50 mM Tris[pH 12.45]). Tubes were gently inverted until the suspensionpartially cleared (5 to 30 s). After the pH was lowered by theaddition of 40 ,ul of 2 M Tris hydrochloride (pH 7.0), 120 ,ulof 5 M NaCl was added, and the mixture was gently inverteduntil a thick white precipitate formed. Afterwards the lysatewas kept on ice for 1 to 3 h before being centrifuged in amicrofuge B (Beckman Instruments, Inc., Palo Alto, Calif.)for 30 min at 4°C. The supernatant was decanted into a cleantube and extracted with 0.6 ml of phenol (equilibrated withan equal volume of 0.5 M NaCl). After centrifugation for 10min at 4°C, the top aqueous layer was carefully transferredto a clean microfuge tube. An equal volume of cold (-20°C)absolute ethanol was added, and the DNA was precipitatedduring overnight storage at -20°C. This DNA was collectedby centrifugation for 30 min at 4°C and was suspended in 10,ul of TE buffer (50 mM Tris, 20 mM EDTA [pH 8.0]). Thecontaminating RNA was digested by adding 2 ,ul of RNase (1mg/ml, preboiled for 10 min) and incubating the mixture for30 min at 37°C. The RNase-treated sample was stored at 4°C.

Plasmids from E. coli were isolated by the method of Kadoand Liu (14).

Gel electrophoresis. Test samples in TE buffer were pre-pared for electrophoresis (14) by adding 2 ,ul of loadingbuffer, i.e., 0.25% bromcresol purple and 50% glycerol(vol/vol) in 50 mM Tris acetate electrophoresis buffer (40mM Tris, 20 mM acetic acid, and 2 mM disodium EDTAbrought to pH 7.9 with glacial acetic acid). Electrophoresis,in a horizontal slab gel of 1% agarose (SeaKem, ME; FMCCorp., Marine Colloids Div., Rockland, Maine) was for 2 hat 7 V/cm (Heathkit model IP-17 power supply; Heath Co.,Benton Harbor, Mich.). Gels were stained for 30 min in 200ml of Tris acetate buffer with 0.5 ,ug of ethidium bromide perml and then destained for 5 min in running distilled water.Polaroid photographs were taken with a transilluminator(model 3-3000; Photodyne, New Berlin, Wis.), type 57instant black-and-white film, and a Wratten 23A filter.

Since open-circle (oc) and linear (lin) forms of smaller(<10 MDa) plasmids move more slowly through mostagarose gels than their intact covalently closed-circle (ccc)counterparts, multiple bands in agarose gels can representdifferent forms of a single plasmid (1). The difficulty indeciphering multiple bands in a gel was resolved byreelectrophoresing the gel lanes in a direction perpendicularto the first run (10). Lanes cut from the stained first gel wereplaced in a fresh gel and irradiated by holding a shortwaveUV light (Mineralight UVSL-25; Ultra-Violet Products,Inc., San Gabriel, Calif.) 5 to 10 cm from the gel for 30s. TheUV irradiation nicks the ccc plasmid into oc form so that inthe second electrophoresis the UV-nicked form has the samemobility as the corresponding naturally nicked form in thefirst run. The procedure also unmasks ccc plasmid that mighthave comigrated with a nicked oc form of a different plasmidor with chromosomal DNA.

Molecular sizes were determined from a computer-generated parabolic standard curve based on E. coli plas-mids of 80, 35.8, 3.7, 3.4, 2.6. 2.0. 1.8, and 1.4 MDa.Clostridium plasmid sizes reported are mostly the averagesof at least six determinations.

Curing procedures. When 3- to 6-week-old colonies fromBHI or GYPT agar plates were streaked onto fresh medium,colonies of different morphologies were often observed.Representatives of these and of normal-appearing colonieswere subcultured, and their toxigenicity and plasmid contentwere determined.

Direct curing was by inoculating toxic strains into GYPTbroth containing 0.001% DOC, incubating the culture for 24h at 35°C, and plating dilutions of the culture on GYPT agar(25). Subcultures for further testing were made from coloniesboth with normal-appearing and with altered morphology.The three toxic cultures used for curing studies were (i)strain A73, chosen because its colonies gave the clearestresponse in the immunodiffusion toxicity test (see below),(ii) strain 62A, chosen because its plasmids had been previ-ously examined (20; Kahn and Pierson, unpublished results),and (iii) strain 90A, chosen because it gave the highestfrequency of nontoxigenic variants.Immunodiffusion test for toxigenicity. Isolates were

screened for toxigenicity by an antitoxin overlay immuno-diffusion method (5, 6). Antitoxin was the serum of a rabbitimmunized with the toxoid made by treating a solution oftype A toxic crystals with Formalin. If the culture was usedfor plasmid studies, the immunodiffusion result was con-firmed by testing the toxicity of a 4- to 5-day-old BHI culturein mice (0.5 ml intraperitoneally). For the immunodiffusionassay, only clear rings of immunoprecipitate were counted.Zones of haziness were ignored.

RESULTS

Plasmid isolation. In agreement with other reports (23;Kahn and Pierson, unpublished results), one of the diffi-culties we encountered in isolating plasmid DNA from C.botulinum was the lysing of cells. Exponentially growingcells harvested 8 h after inoculation were easiest to lyse.Plasmid yield was noticeably lower if cells came fromcultures incubated for 12 h, and virtually no plasmid DNAwas recovered if cells were from cultures incubated for morethan 14 h.Lysozyme plus the SDS solution was sufficient for lysis of

cells from 8-h cultures, but slightly better lysis was obtainedwhen TBL-1, a bacteriolytic enzyme, was included. C.botulinum lysates were never completely cleared after addi-tion of the SDS solution, though C. sporogenes lysatesalways cleared completely.A second difficulty encountered in plasmid isolation in

neutralizing DNases (23; Kahn and Pierson, unpublishedresults). Diethyl pyrocarbonate, which neutralizes Clostrid-ium perfringens DNases (2), was not effective in the presentwork. Heating at 55°C for 10 min, a high concentration ofEDTA (up to 200 mM), or an alkaline SDS solution was alsonot completely satisfactory, since plasmid DNA was oftennicked during recovery.The requirement of only 5 ml of culture for plasmid

recovery instead of 100 ml or more, as needed in previousstudies (20, 23; Kahn and Pierson, unpublished results),simplified the examination of numerous strains and variants.However, the chromosomal DNA is not completely re-moved, and a slight background fluorescence is present insome gels.

Immunodiffusion. Isolates which were toxigenic for micewere tested in the immunodiffusion assay. To determine theoptimum assay conditions, different antitoxin concentrationswere tested, and the plates were read at different times afterstaining. The size and intensity of the immunoprecipitatehalo varied with the strain, but 0.3 IU of antitoxin per ml ofoverlay agar was chosen as the working concentrationbecause it allowed the earliest correct identification afterstaining of all toxigenic cultures (Table 1). When this anti-toxin amount was used, halos around toxigenic colonieswere clearest 8 to 24 h after staining, and no immunoprecip-

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54 WEICKERT ET AL.

TABLE 1. Effects of antitoxin concentration in the overlay agarand time after staining plates on the percentage of 18 mouse toxic

cultures producing immunodiffusion rings% Cultures producing immunodiffusion rings after treatment

Timea of with antitoxin (IU/ml)observation

0.1 0.2 0.3 0.4 0.5 1.0 1.5 2.0

Immediate 0 0 61 39 67 6 11 61 day 0 67 100 83 NTb NT NT NT3 days 11 50 89 72 100 100 89 67

a After staining.bNT, Not tested.

itate rings were produced by variants which were nontoxicby the mouse test or by C. sporogenes cultures.

Plasmids in wild-type C. botulinum. Gel electrophoresis ofplasmid DNA from type A strains showed up to six bands.Five bands migrated faster and one more slowly than chro-mosomal DNA. Two-dimensional gel electrophoresisshowed that three of these bands were different forms of thetwo smaller plasmids (Fig. 1). The molecular sizes of thethree unique ccc plasmids were 4.3, 6.8, and 36 MDa (Fig. 2,lane A73). The 36-MDa band is not seen in Fig. 1 becauserapid degradation of this band occurred when samples werestored at 4°C before electrophoresis.The 4.3-MDa ccc plasmid migrated with an apparent

molecular size of 10.5 MDa when it was in the oc form. The6.8-MDa ccc plasmid migrated with an apparent size of 17.1MDa when in the oc form and an apparent size of 12.1 MDawhen in the lin form. An oc or lin counterpart of the 36-MDaplasmid was not present.A slowly moving band indicative of a 61-MDa entity was

occasionally encountered. It was found at least once in eachof four parent strains and two variants, but was not alwaysfound when the strains were retested.

v8 0v3 6

_ e ~~~~~~~~~~~ch r

FIG. 1. Plasmid bands of three C. botulinum strains after single-and two-dimensional (2d) agarose gel electrophoresis. Strain A73(A) has five bands, including the oc and lin forms of the 6.8-MDaplasmid and the oc form of the 4.3-MDa plasmid. Strain 62A (B) isidentical to strain A73, except it lacks the 6.8-MDa lin form. StrainA3 (C) has only the three forms of the 6.8-MDa plasmid. The 2dlanes were electrophoresed first from left to right and then from topto bottom of the gel and have the oc bands migrating above (moreslowly than) the diagonal of fluorescence. The oc forms in 2d gelsmigrate at rates identical to those of the original bands and directlyabove their ccc plasmid counterparts. D and E are E. coli referenceplasmids of 36, 3.7, 3.4, 2.6, 2.0, 1.8, 1.4 (all in lane D), and 80 MDa(E). The nicked 80-MDa plasmid (E 2d) does not migrate into the 2dgel. Numbers on the right are selected reference plasmid sizes inMDa. chr, Chromosomal DNA.

804

ch r-ec

_1 ~ ~~~~ ~~~~~~~~~~~~~ncc

3.74 )

FIG. 2. Agarose gel of plasmids from DOC-treated C. botulinumtype A strains. Lanes show the following: A and B, size referencemarkers from E. coli; C and D, nontoxigenic strain A73 variants; Gand H, toxic variants of strain A73; E and F, nontoxigenic variantsof strain 90A; and I and J, toxic strain 90A plasmids; A73, plasmidprofile of toxic strain A73. Numbers on the left are molecular sizesin MDa of selected standard bands. chr, Chromosomal DNA.

The ccc plasmids of the wild-type C. botulinum culturesare summarized in Table 2. If the 61-MDa entity is notincluded, three plasmids were the maximum found in anyone strain. No plasmid was found in strains 69A and 11A.

TABLE 2. Plasmids of C. botulinum type A and C. sporogenesand their correlation with toxigenicity in type A strains and their

variants

Presencea of Plasmids of size (MDa):Species and strain Toxicity 4.3 5.6 6.8 26 36 61

C. botulinum type A62A + + + - +62A-V1Db + - + - +62A-V2D - - - - -62A-V3D - + - - -62A-V4D + - - - -A73 + + + - +A73-V1D + - + - +A73-V2D + - + - -A73-V3D - + - (+) -90A + - + - +90A-VlD + - - - -90A-V2D - - - (+) -A3 + - +A3-V1Sc + - - - -A3-V2S - - - - -56A + + + - +56A-V1S + - + - +19A + + - -19A-VlS - + - - -Hl + - + - +172A + - + - - -

A109 + - + - + -

11A + - - -

69A + - - - -Af84-SC2 + - - - -

C. sporogenes4411 - + - -4447 - - - -4449 - - - -4452 - + - +4472 - - + -4475 - - - -PA3679-66 - - + +a (+), Not recovered consistently.b Variant from DOC-treated culture.C Variant arising spontaneously.

APPL. ENVIRON. MICROBIOL.

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C. BOTULINUM PLASMIDS AND TOXICITY 55

The single plasmid of strain 19A and that of strain 3A weredifferent in size. There was no plasmid common to all thetype A strains tested.

Variants of C. botulinum. Colonies of nontoxigenic vari-ants of C. botulinum type A tended to be round andtranslucent, whereas toxigenic colonies were generallyrough, irregular, and opaque. There were numerous excep-tions, however, and colony morphology was not an adequatepredictor of toxigenicity. From 5 to 12% of the coloniesobtained by the DOC curing method were nontoxigenic bythe immunodiffusion test.

Plasmids in isolates from DOC-treated cultures of strainsA73 and 90A are illustrated in Fig. 2. Two toxigenic isolatesof A73 (lanes G and H) lacked the 4.3-MDa ccc plasmid ofthe parent culture (lane A73). Two nontoxigenic isolates hadthe 4.3-MDa but not the 6.8- and and 36-MDa ccc plasmidsof the parent, but they also had a 61- and a 26-MDa band(ust above the chromosomal DNA band in lanes C and D),neither of which had been observed in the parent strain.Some nontoxigenic isolates of strain 90A obtained with

DOC had only the 26- and 61-MDa bands (Fig. 2, lanes E andF). The different plasmid profiles of strain 62A isolates aresummarized in Table 2, along with those of variants ofstrains 90A and A73.

Plasmids of C. sporogenes. Three different-sized ccc plas-mids were found in the seven strains of C. sporogenestested. Two were of the same size (4.3 and 36 MDa) asplasmids found in some C. botulinum strains; the third was a5.6-MDa plasmid not present in the toxigenic species. Nostrain had all three plasmids, and three strains had none(Table 2). The 6.8-MDa plasmid of the toxigenic species wasnot seen in C. sporogenes.

DISCUSSION

Previously reported methods for isolating C. botulinumplasmids used no less than 100 ml of culture (20, 23; Kahnand Pierson, unpublished results), whereas the presentmethod requires only 5 ml. Using this method of plasmidextraction, we observed that the three intact (ccc) C. botu-linum type A plasmids most prevalent were 4.3, 6.8, and 36MDa in size. Presumably these plasmids correspond to the3.6-, 6.2- to 6.9-, and 33.5-MDa plasmids reported by others(20, 23; Kahn and Pierson, unpublished results). A plasmidof 15 to 18.9 MDa has been reported (20, 23; Kahn andPierson, unpublished results), but we were unable to confirmthat observation although the oc form of the 6.8-MDa cccplasmid has the mobility corresponding to that of a17.1-MDa ccc plasmid.A DNA band of about 61 MDa was isolated at least once

from four parent cultures and two variants of these cultures.It is probably not the oc or dimeric form of the 36-MDaplasmid, since these do not enter the 1% agarose gels used inour study. Also, the observation of the 61-MDa band inseveral preparations lacking the 36-MDa plasmid suggeststhat these two may be distinct plasmids. There is also thechance that the 61-MDa unit represents prophage DNA,since phages are common in C. botulinum strains (25).A unit of about 6 MDa appeared in some preparations of

DOC-induced variants of-two type A strains which had beenpartially cured of plasmids. If this unit were present in parentstrains but masked by contaminating chromosomal DNA, itshould have been revealed during two-dimensional electro-phoresis, but was not. The origin of this plasmid is unknown.One possibility is that the DOC treatment results in thedeletion of a portion of the 36-MDa plasmid, yielding the

26-MDa band. Hybridization analysis could confirm thispossibility.The comigration of two of the three C. sporogenes plas-

mids with two plasmids from C. botulinum type A strains isin agreement with the view that the two species are geneti-cally related (16, 18, 27). A previous study (Kahn andPierson, unpublished results) found one or more plasmids ofapproximately 10, 25, or 40 MDa in three of six strainsexamined. We also found plasmids in about half (4 of 7) ofthe C. sporogenes strains we examined. However, in an-other study (23), no plasmid DNA was found in any of thenine C. sporogenes strains examined, including four havingthe same designations as four of those used in the presentstudy. In three of these four strains, we found no plasmids,either, but in strain 4472, a plasmid of 5.6 MDa wasobserved. This difference may be due to different sources forthe strains, plasmid loss from some strains, or an inappro-priate plasmid isolation method being employed in the otherstudy.

Toxicity of C. botulinum is normally assayed by growingthe strain to be tested in a liquid medium and showing thatthe culture fluid is lethal to mice by the intraperitonealinjection route. The method is not well suited for examiningmany cultures because of the expense and difficulties ofusing mice. A variation of the immunodiffusion methoddevised by Ferreira et al. (5, 6), in which antitoxin wasincorporated only in the agar overlay, proved satisfactoryfor a strain screening program. It provided a 100% accurateidentification of toxigenic colonies.

In curing the strains of toxicity, we initially relied oncolony morphology changes as an indicator of potentialnontoxigenic derivatives. Takumi et al. (25) reported thatnontoxigenic colonies of type A strain 190L were large,opaque, and white. Though we did observe different colonymorphologies after plating broth cultures treated with deter-gents, opaque white colonies were infrequent, and in somestrains, the normal colony was large, opaque, and white.Spontaneous variants of unusual colony morphology werealso screened for toxicity by the immunodiffusion test or forplasmids or for both. We found that colony morphology wasnot a reliable indicator of toxigenicity. Because of thisfinding, we tested both normal and unusual colony types bythe immunodiffusion assay to identify the nontoxigenic vari-ants.Though a frequency of generation of nontoxigenic variants

for strain A19OL of up to 94.4% was reported (25), ourfrequency of curing was not so spectacular (only 5 to 12%).This difference could be due to differences in the strains usedor to other unidentified factors.

If strain Af is assumed to be type A, then plasmids werefound in 9 of 12 toxigenic type A strains, a frequency notsignificantly different from the 12 of 15 strains previouslyreported (20, 23; Kahn and Pierson, unpublished results).Since approximately 22% of the strains examined so far haveno observable plasmids, it seems unlikely that plasmids areresponsible for toxin production unless a system such as thatobserved in Bacillus thuringiensis exists. Genes for B.thuringiensisc-endotoxin were found both on a plasmid andin the chromosome in B. thuringiensis subsp. kurstaki (9),while the c-endotoxin genes in B. thuringiensis subsp.thuringiensis, B. thuringiensis subsp. alesti, B. thuringiensissubsp. galleriae (8), and B. thuringiensis subsp. israelensis(26) are uniquely plasmid borne.

Toxicity in the type A strains we examined does notappear to be plasmid directed unless a plasmid other thanthose we have been able to recover is responsible. Even the

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larger plasmid we infrequently isolated was found innontoxigenic variants and therefore is unlikely to carry thetoxin determinant. The genetics of type A toxin productionremains unknown, as does the function of the plasmids, butour results do show that toxin production is not related to theobserved plasmids in a direct manner

ACKNOWLEDGMENTSWe thank Peter A Kahn and Merle D. Pierson for making available

their plasmid isolation method.This research was supported by the College of Agricultural and

Life Sciences, University of Wisconsin, Madison, and by contribu-tions to the Food Research Institute, University of Wisconsin, bymember industries. M.J.W. was supported by Public Health Servicetraining grant no. 5 T32 GM-7133-10 from the U.S. Department ofHealth and Human Services.

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