Project Abstract / Adhesion-Mediated Enrichment of Purple Sulfur Bacteria

Carol BarfordMicrobial Diversity

Summer 1993

An independent research project was conducted which investigated the effects of adhesion to

different solid substrata on bacterial enrichments from an environmental inoculum. Stainless

steel, flint glass, polystyrene and tissue-culture polystyrene were chosen for their ready

availability and differing surface wettabilities, a characteristic which is known to influence

biological adhesion. Purple sulfur bacteria were enriched from samples of the pink layer of

Great Sippewisset Marsh microbial mat in anaerobic cultures to which the solid substrata were

added as 0.5 x 8.0 cm strips. The solid substrata were passaged through three stages of

enrichment medium. At each stage the culture supernatants and the attached microbial

community were examined by phase contrast microscopy. At the first and third stages

subsamples of the solid strips were taken for bacteriochiorophyll a (Bchl a) determination and

preparation for scanning electron microscopy. Bchl a content was also determined for the

third stage supernatants (bulk phase). Polystyrene appeared to be a favorable substratum for

the adhesion and growth of a Chromarium strain. By the third stage of enrichment a relatively

dense, homogeneous biofllm of a putative Chromarium had developed on polystyrene, which

was one of the less wettable substrates. This biofilm contained high amounts of Bchl a

(normalized against protein content), perhaps due to light attenuation by the thick biofilm.

Bacterial attachment to other substrata was less dense and resulted in mixed-morphotype

aggregates. In all cases the final bacterial enrichment in the bulk phase differed from the

biofilm.

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Adhesion-Mediated Enrichment of Purple Sulfur Bacteria

Carol BarfordMicrobial Diversity

Summer 1993

Introduction

Adhesion of bacteria to solid surfaces is important in many natural microenvironments,

including soil crumbs, suspended particulate organic matter, and intra-animal surfaces

(Costerton et al. 1985). Physiology, morphology and community structure can differ between

attached and bulk-phase bacteria (Fletcher 1985). Such differences have practical implications

for clinical medicine, e.g. biofilm formation on implanted materials; water delivery and

wastewater treatment systems, and many other environments (Characklis and Marshall 1990).

Bacterial adhesion is a complex phenomenon which depends on surface characteristics

of the solid substrate and the bacterial cell, as well as environmental parameters such as pH

(Marshall 1976). Because it depends on several physical and chemical factors, adhesion

behavior is difficult to model. However, crude indices of surface characteristics can be used

for preliminary correlation with biological adhesion (Gerhart et a]. 1992). In this project solid

substrata with different wettabilities were incubated with environmental inocula under

conditions known to enrich for purple sulfur bacteria. It was hypothesized that bacterial

adhesion to the substrata would result in enrichments of different purple sulfur bacterial

communities after a few passages through the selective medium. Purple sulfur bacteria were

chosen for enrichment because of their growth habit in Great Sippewisset Marsh, where they

are present as a distinct, cohesive layer of pink sediment. This suggests the ability to adhere

and constitutes a targeted inoculum (Gibson et al, 1984).

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Materials and MethodsCSolid substrata with a range of surface wettabilities were chosen. Table I lists

published values for wettability as well as measured values obtained by the same procedure

(Gerhart et al. 1992).

Table 1. Surface Wettabilities

Surface Wettability Wettability

(measured) (published)Stainless Steel 20.8

Flint Glass 68.9 95.0* borosilicate glassPolystyrene 20.5 25.6Tissue Culture Polystyrene 30.9 46.3

Solid substrata were cut into 0.5cm x 8cm strips and cleaned by 2 washes with 70% ethanol

and one wash with distilled water before inoculation. The inoculum was prepared from the

pink sediment layer of Sippewisset Marsh mat by combining approximately equal volumes of

sediment and sterile seawater in a screwcap tube with one drop of 1 M Na2S and vortexing.

Two ml of the resulting suspension were added to each of four 50 ml Pfennig bottles

containing 1) Pfennig’s purple sulfur medium (marine) with sodium acetate added to a final

concentration of 5mM (Pfennig and Truper 1992) and 2) each of the four substrate strips (four

bottles, sixteen strips). The enrichments were incubated in bright incandescent light at room

temperature for 5 days, which was sufficient time for the strips to be heavily colonized. After

the first incubation,every strip from three bottles was gently washed and subsampled for 1)

bacteriochlorophyll a (BehI a) determination , 2) preparation for scanning electron microscopy

(SEM), and 3) transfer to sterile purple sulfur medium containing a clean strip of the same

material. Two more iterations of the subsampling and transfer were made at 5 day intervals.

However SUM and Bchl a analyses were not performed at the second enrichment stage. At

the third enrichment stage Bchl a was also determined for the supernatants. The fourth bottle

at each stage of the enrichment was used for phase microscopic examination of the culture

supematants and attached bacteria. Bchl a was determined by A of acetone extracts and

normalized against total cell protein, which was determined by a modified Folin assay.

Results

1. Phase microscopy. In the first enrichment stage, heterogeneous microbial aggregates

formed on each solid substrate. These included purple sulfur bacteria and cyanobacteria,

spirochaetes, ciliates, diatoms and unidentified rods and spirillae. Attached microbial biomass

was visibly higher on the upper side of all strips, perhaps due to greater light availability.

The polystyrene strips supported relatively large, brownish-purple aggregates of Chromatiurnn(possibly C. vinosum). The second enrichment stage was more homogeneous and contained

few eukaryotes.. It was dominated by Chromarium and Thiocapsa strains, which formed very

homogeneous flocs in the bulk phase. At the second stage it became clear that the attached

and bulk phase bacterial communities were qualitatively dissimilar. At the third stage,

colonization of the substrata was slower, and few cells were attached to stainless steel after 5

days. The bulk phase of culture bottles was dominated by Thiocapsa (possibly T.

roseopersicina) with many smaller highly motile rods also present, while the solid substrata

were covered with finer-grained patterns of Chromatium.

2. SUM. In the first enrichment series, microbial colonization and growth on 1-2 cm length

subsamples of solid substrata was heterogeneous enough to preclude the possibility of finding

“representative” fields of view at magnifications higher than about 2,000x. However, SUM

nfl.., -4

enabled visualization of general patterns. Colonization of steel and glass was relatively fine

grained with mixed-morphotype aggregates, while aggregates on polystyrene and tissue-culture

polystyrene (TCPS) were larger and composed of single morphotypes. In the third enrichment

stage glass and polystyrene strips were covered with quite dense, homogeneous films of

Chromarium with easily visible capsular exopolysaccharide. TCPS was covered with a less

continuous biofilm of Chromatium, and steel strips exhibited patchy colonization by several

morphotypes.

3. Bchl a analysis. The results of Bchl a analyses on solid substrata and supematants are

given in Table 2. Normalized Bchl a content of attached bacteria was consistently highest for

polystyrene but no significant differences by substrate were detectable in the third stage

supernatants.

Table 2. Bchl a normalized by protein content (mg Bchl a/g protein).

. Stainless Steel Flint Glass Polystyrene TCPS

1st stage strips 38.1 ± 7.1 38.5 ± 2.5 135 ± 62 49•g*

3rd stage strips protein too low 27.3 ± 2.1 199 ± 43 58.1 ± 28

3rd stage spnt. 18.5 ± 8.9 33.5 ± 18 5.1* 42.3*

* single datum

Conclusions

It appears that polystyrene may be used to enrich for putative Chromatium (vinosum 7)

from a Sippewisset Marsh mat inoculum. Biofllms on polystyrene were homogeneous and

C) relatively dense as visualized with SEM. The enrichments attached to polystyrene differed

from the suspended bacterial community which suggests that aggregates and films found on

polystyrene grow in situ from a few attached cells rather than develop in an attachment-

detachment equilibrium with the bulk phase. This conclusion is also supported by the

homogeneity of bacterial aggregates on polystyrene from the first enrichment stage and the

amount of exopolysaccharide evident in the third stage biofllm SEM’s. Bchl a analysis

suggest that putative Chromatium cells attached to polystyrene may respond to light

attenuation by the thick biofilm by producing more Bchl a. The results suggest that this is a

quantitative response to bioftim thickness rather than a qualitative characteristic of the

attached putative Chromatiwn because the biofilms on glass were also homogeneous but did

not contain as much Bchl a per protein.

References

Characklis, W. and K. Marshall (eds). 1990. Biofilms. John Wiley and Sons, New York.

Costerton, J., T. Marrie and K. Cheng. 1985. Phenomena of bacterial adhesion. pp 3-43. In0. Savage and M. Fletcher (eds), Bacterial Adhesion. John Wiley and Sons, New York.

Gerhart, D., D. Rittschof, I. Hooper, K. Eisenman, A. Meyer, K Baler and C. Young. 1992.Rapid and inexpensive quantification of the combined polar components of surfacewettability: application to biofouling. Biofouling 5: 251-259.

Gibson, 0., E. Leadbetter and H. Jannasch. 1984. Great Sippewisset Marsh: A summary ofprojects carried out by students in the Microbial Ecology Course of the Marine BiologicalLaboratory, Woods Hole, During Summers 1972-1981. pp. 95-100. In Y. Cohen, R.Castenholz and H. Halvorson (eds), Microbial Mats: Stromatolites. Alan It Liss, New York.

Fletcher, M. 1985. Effect of solid surfaces on the activity of attached bacteria. In 0. Savageand M. Fletcher (eds), Bacterial Adhesion. John Wiley and Sons, New York.Marshall, K. 1976. Interfaces in Microbial Ecology. Harvard University Press, Cambridge,MA. 156 pp.

Pfennig, N. and H. Truper. 1992. The Family Chromatiaceae. pp. 3200-3222. In A. Balows,H. Truper, M. Oworkin, W. Harder and K. Schleifer (eds), The Prokaryotes. Vol. 4, SecondEdition. Springer-Verlag, Berlin.

0 Zisman, W. 1964. Relation of the equilibrium contact angle to liquid and solid constitution,

pS-fr 6

pp.1-51. In it Gould (ed),. Contact Angle, Wettability and Cohesion. American ChemicalSociety, Washington, D.C.

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Project Abstract / Effect of Culture Conditions on Exopolysaccharide Production

Carol BarfordMicrobial Diversity

Summer 1993

Exopolysaccharide production in an enrichment of purple sulfur bacteria was studied.

A pure culture of putative Thiocapsa roseopersicina was isolated from an enrichment from

Great Sippewisset Marsh samples, grown in selective medium, then inoculated into

experimental media in a factorial matrix of three environmental parameters. The parameters

were pH (6.8. 7.2, 7.8), light intensity (200 and 20 uEIm2), and salinity (freshwater and

marine. Following 5 days’ incubation the cultures were subsanipled for total polysaccharide,

protein, and bacteriochiorophyll a content. Due to the short growth interval, protein was too

low to be determined in many of the cultures. Thus, no statistically significant relationships

between environmental parameters and exopolysaccharide (BPS) production were observed.

However, generally higher BPS production occurred at pH 7.8, which may indicate stress on

the bacterial isolate at that pH.

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flThe Effects of Selected Environmental Parameters on Exopolysaccharide Production by

a Purple Sulfur Bacterium

Carol BarfordMicrobial Diversity

Summer 1993

Introduction

In many environments, bacteria live in aggregates or biofilms which consist largely of

cells in matrices of exopolysaccharide (e.g. Foster 1981, Costerton et al. 1985). The capsular

and peripheral exopolysaccharide (EPS) create protective microenvironments for bacteria.

BPS has been found to protect bacteria from desiccation (Roberson and Firestone 1992), metal

and antibiotic toxicity, shear stress, and starvation conditions (Ralph Mitchell, personal

communication). Conversely, BPS production can increase during physiological stress such as

desiccation (Roberson and Firestone 1992). In this project exopolysaccharide production by a

C) putative Thiocapsa roseopersicina strain was measured following incubation under different

pH, light, and salinity conditions in anaerobic liquid medium. It was hypothesized that

conditions differing from the selective enrichment conditions by which the strain was isolated

would induce BPS production. Salinity, light and pH were chosen because of the ease of

manipulation of these variables over relatively short incubation periods. Purple sulfur bacteria

were chosen as experimental organisms due to apparent BPS production in their growth habits

as aggregates and microbial mats.

Materials and Methods

A pure culture of putative Thiocapsa roseopersicina was isolated from an agar shake

series kindly provided by Minoru Wada, a fellow student. The original enrichment inoculum

was collected from Great Sippewisset Marsh. A cohesive, maroon-colored colony was

0 picked from the agar and tranferred to selective purple sulfur medium (Pfennig and Truper

a _ •.Z.

1992), amended sodium acetate to 5 mM final concentration and marine salts solution. The

culture was grown in bright incandescent light at room temperature for 8 days, then

transferred to flesh medium (backup culture) and inoculated into the experimental media. At

this point BPS could be seen in negative-stained wet mounts by brightfield microscopy. The

experimental culture conditions included duplicates of all factorial combinations of three pH

levels (6.8, 7.2, 7.8), two light levels (200 and 20 uEIm2), and two salinities (freshwater and

marine as defined by Pfennig and Truper, 1992). Experimental cultures were incubated for S

days. At the end of incubation the tubes were vortexed at high speed for 5 minutes to

disrupt the capsular EPS, then centifuged for 8 minutes at 14,000 rpm. The supernatants were

reserved for saccharide determination by the anthrone reaction. This was done by adding 2

ml of anthrone reagent (2 mg/nil in concentrated H2S04)to I ml of supematant, boiling the

mixture for 15 minutes, cooling on ice and measuring A6. The pellet was extracted with

100% acetone for 5 hours in the dark, and the supernatants used for determination of

bacteriochlorophyll a (Bchl a) by measuring A770. The protein content of the pellet was

determined by the Folin assay. BPS production in the cultures was normalized by protein

content and normalized Bchl a content was intended to be an index of light level adaptation.

Results

The results of all determinations are given in Table 1. There were no statistically

different levels of BPS production or Bchl a content between experimental treatments.

Conclusions

Although no statistically significant differences were found, possible trends can be

inferred from the data. In general there was higher normalized BPS production at pH 7.8.

This could indicate low tolerance of high pH in this putative Thiocapsa strain. There was

also a trend of higher protein and normalized Bchl a production, but not EPS production, at

high light level. This could be interpreted as reduced BPS production under favorable growth

conditions. The paucity of data resulted from insufficient incubation time. The Folin protein

assay appeared to be the least sensitive of the three analyses, and because protein content was

used to normalize the other data undetectably low protein values rendered the other values

useless. However, it would be easy to improve upon this project by increasing incubation

time, or possibly by adding solid substrata as incubation variables.

References

Costerton J., T. Marrie and K. Cheng. 1985. Phenomena of bacterial adhesion. pp.3-43. In 0.Savage and M. Fletcher (eds), Bacterial Adhesion. John Wiley and Sons, New York.

Foster, R 1981. Polysaccharides in soil fabrics. Science 214: 665-667.

Pfennig, N, and H. Truper. 1992. The Family Chromatiaceae. pp. 3200-3222. In A. Balows,H. Truper, M. Dworkin, W. Harder and K. Schleifer (eds), The Prokaryotes. Vol. 4, SecondEdition. Springer-Verlag, Berlin.

Roberson, E. arid M. Firestone, 1992. Relationship between desiccation and exopolysaccharideproduction in a soil Pseudomanas sp. Applied and Environmental Microbiology 58: 1284-1291

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_,

* high light = 200 tiE/rn2, low light = 20 uE/m2** Freshwater and marine as defines by Pfennig and Truper (1992).

Units = mg glucose equivalents per mg protein*44* Units = mg BehI a per g protein# indicates single datum, “und” indicates undetectably low levels.

C) .Table 1. EPS (glucose equivalents) and Echi a contents of Thiocapscz cultures.

pH light4 salinity4 BPS444 Bchl a4446.8 high fresh 2.02* 17.4 ± 6.96.8 high marine 0.0618* 13.3 ± 3.66.8 low fresh O.244# 19.7*6.8 low marine und 14.5*7.2 high fresh 0.063* 18.3 ± 147.2 high marine 0.183 ± 0.08 16.3 ± 157.2 low fresh und und7.2 low marine und und7.8 high fresh 2.90* 22.6*7.8 high marine 0.895* 13.9 ± 3.17.8 low fresh und und7.8 low marine 4.00 ± 2.8 und

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