Electron Microscopic Study of a Slime Layer

Transcription

1 JOURNAL OF BACTERIOLOGY, July 1969, p American Society for Microbiology Vol. 99, No. I Printed in U.S.A. Electron Microscopic Study of a Slime Layer H. C. JONES, I. L. ROTH, AND W. M. SANDERS, III Department of Microbiology, University of Georgia, Athens, Georgia 30601, and Southeast Water Laboratory U.S. Department of the Interior, Athens, Georgia Received for publication 23 April 1969 Slime layers are being studied in our laboratories in an attempt to understand their functions in the control of pollution in natural streams. A method for fixing, staining, and embedding microorganisms in the intact slime has been developed. In this method, epoxy resin discs are placed in a holder and are introduced into a simulated stream. After various periods of time the discs are punched out of the holder into the fixative. The disc with the attached slime is fixed, stained (4% osmium tetroxide plus ruthenium red), dehydrated, and embedded in epoxy resin so that thin sections can be cut through the vertical plane of the slime mass. Such thin sections permit detailed examination of the attached layer, the surface-slime interface, the spatial relationships between cells in the vertical slime structure, and the strands of extracellular material between and around cells. No special attachment structures were noted as the cells appeared to be attached to the surface by extracellular material alone. This material was observed in strands and netlike forms between cells which are positioned 1 to 4,m apart in the slime. Factors responsible for the occurrence and growth of bacterial communities in streams are being studied in our laboratories (16, 19). We have developed a technique for fixing, staining, embedding, thin-sectioning, and examining by electron microscopy the organisms in the slime layer. Ultrathin sectioning through the vertical plane of the slime mass, including the attachment surface, enables detailed examination of the primary or attaching layer, the slime-surface interface, the spatial relationships between cells in the vertical slime structure, individual cell morphology, and the slime material between and around the cells. It is anticipated that this method will prove useful in studying the ecology of naturally occurring microbial films in aquatic systems. MATERIALS AND METHODS Apparatus. Preformed epoxy resin discs were fitted into holes drilled into Plexiglas microscope slides which were in turn inserted into a growth chamber. Upon removal from the chamber, these discs with the organisms attached directly to the exposed surface were placed directly into a container of fixative to minimize handling. The discs were formed by filling the cap of a BEEM capsule (Better Products for Electron Microscopy, Inc., Bronx, N.Y.) with either Maraglas 655 (3), Maraglas 732 (2), or Epon 812 (10). After hardening, the caps were stripped away, leaving a relatively smooth surface on one side of the disc. Eight holes, each 9 mm in diameter, were drilled in each Plexiglas slide approximately 0.5 inch (1.27 cm) apart (edge to edge), and the discs were inserted aseptically after the system, slide, and discs had been sterilized separately. Sterilization. Either mercuric chloride (1%) or (Bpropiolactone (6%) was used to disinfect the materials since both Plexiglas and epoxy resins are damaged by heat. The materials were then rinsed overnight in sterile distilled water to rid them of residual disinfectant. Culture conditions. The slides containing the epoxy discs were inserted into continuous-flow growth chambers which were being operated by the Ecological Engineering Activity, Southeast Water Laboratory (16). The cylindrical chambers were approximately 6 inches (15.24 cm) high and 3 inches (7.62 cm) in diameter with a mean volume of 519 ml. The substrate, 20,g of nutrient broth (BBL) per ml, flowed through the chamber at the rate of 5 ml/min, with a mean residence time of 1.9 hr. The temperature of the system at the time of sampling was 35 C, and the radial velocity of the medium within the chamber was maintained at 1.5 ft/sec by a magnetic paddle mechanism. The chambers were seeded with lyophilized microorganisms which had been previously obtained from a polluted stream. The microscope slide, when inserted into the growth chamber, fit snugly against the chamber wall so that the organisms attached onto the exposed surface of both the slide and the epoxy resin discs. After various periods of growth (7 to 9 days), the slide was removed from the growth chamber and the discs were gently pushed into containers of fixative. Fixation and staining. Seven-day samples (Fig. 1 and 2) were fixed with several fixatives or fixative and ruthenium red (11; K & K Laboratories) mixtures with and without prior staining in 0.1% alcian blue (12) as follows. 316

2 VOL. 99, 1969 ELECTRON MICROSCOPIC STUDY OF SLIME LAYER 317 Specimens were stained with alcian blue (0.1%) for 1 min and then were fixed in 2% KMnO4 for 30 min or fixed by the Kellenberger method (9). Other specimens were stained with 0.1% alcian blue for 2 min and then were fixed and stained in ruthenium redglutaraldehyde-os04 by the procedure of Pate and Ordal (13) or by the Pate-Ordal procedure omitting ruthenium red. Specimens were also fixed with all of the aforementioned fixatives without prior staining with 0.1% alcian blue. Nine-day samples (Fig. 3 through 10) were fixed by the Kellenberger method or by the procedure of Pate and Ordal. Thin-section preparations. After fixation, the specimens were rinsed in several changes of distilled water to remove bxcess fixative and were dehydrated by successive use of 30, 50, 70, and 90% ethyl alcoholwater mixtures applied for 20 min each with two final applications of absolute ethyl alcohol. Infiltration followed, and the discs were embedded in Epon 812 by use of the method of Luft (10) and were hardened by being placed in an oven at 60 C for 72 hr. Ultrathin sections were cut on an LKB 4801A Ultratome with glass knives broken at 450 and 50. Sections were collected on uncoated, 300-mesh copper grids. Some sections were poststained with lead citrate (14) for 15 min, followed in some instances by staining with 2% uranyl acetate (18) for 1 hr. All sections were examined in a JEM 6C electron microscope with an accelerating voltage of 80 kv. Deoxyribonuclease treatment. One of the discs from the 9-day samples was immersed in a solution of deoxyribonuclease (20 mg/ml; Sigma Chemical Co.) for 30 min at room temperature before staining with ruthenium red. RESULTS The sections obtained from samples stained with ruthenium red showed staining of the slime material surrounding the bacteria and those areas where the organisms are in direct apposition to the surface. Designated as polysaccharide-like (P), these areas are shown in Fig. 1 through 10. Sections of the 7-day specimen indicate that a mat is present consisting of microorganisms and polysaccharide-like material (Fig. 1 and 2). This mat is approximately level across the top (Fig. 1, unmarked arrow) and varies in height from 5 to 8,um, as determined from measurements made on micrographs. The epoxy resin surface (AS), to which slime is attached, is visible at the bottom of the mat (Fig. 1 and 2). These micrographs represent the general physical arrangement of the slime layer of the 7-day sample, which differs markedly from the layer as it appears in the 9-day sample. In the 7-day sample (Fig. 1 and 2), the homogenousdistribution of polysaccharide strands is evident. In contrast, the strand distribution in the 9-day sample (Fig. 3) generally shows the slime material compacted in areas adjacent to the microorganisms. In Fig. 3, 4, 7, and 8, a clear, nonstaining or void area (V) is seen surrounding most cells. This void area is enclosed by strands of densely stained polysaccharide-like material. Some cells in the 9-day samples show evidence of plasmolysis (Fig. 4). In Fig. 7, one cell is seen to be surrounded by polysaccharide, whereas another cell adjacent to it shows no extracellular polysaccharide. The cell which is not surrounded with polysaccharide appears to have a bleb forming from the cell wall (unmarked arrow). The cell wall and cytoplasm of this microorganism are stained more densely than the other cell which has the concentric strands of compacted polysaccharide around it. In Fig. 5 and 6, the strands of polysaccharide are homogenously distributed and appear to be emanating from the surface of the microorganisms, whereas in Fig. 4 and 8 the strands of polysaccharide are compacted adjacent to the cell but separated from the cell by a void area. The slime-surface interface of the 9-day samples is illustrated in Fig. 9 and 10; also shown are the microorganisms attached to the surface via densely staining polysaccharide-like material which is identical in appearance to the polysaccharide found between cells. Those samples treated with deoxyribonuclease demonstrated that the polysaccharide-like material surrounding the cells was not destroyed and maintained its threadlike appearance, indicating that it was not composed of deoxyribonucleic acid. Treatment of the slime layer with alcian blue before embedding failed to reveal any extracellular material. When compared with other sections, the samples stained with alcian blue showed cells distorted to such an extent that no micrographs were taken. DISCUSSION The colonization of artificial bare areas presents a vehicle by which many studies of microorganisms grown in nature may be carried out in regard to speciation, succession, and relationships to one another in the community (1). One of the many shortcomings of microbial ecology is the virtual nonexistence of methods by which mixed cultures of microorganisms may be studied. The many problems inherent in such a system are too numerous to list here. In this paper, we attempt to show some physical parameters found in this type of system with emphasis on attachment of these microorganisms to a surface and physical attachment of cells to each other. This was done in the

3 318 JONES, ROTH, AND SANDERS J. BACTERIOL. I/X - AC.0or-~~~~~~~~~~~~~~A -.s,m. A'' L "'* '' 5'" A 4-v 44l FIG. 1. Electron micrograph ofa 7-day sample without poststain. The top of the slime layer is evident (unmarked arrow) as is the artificial surfjce (AS) at the bottom of the slime material. Note the microorganisms (M) embedded in the slime material. X 22,000. FIG. 2. Electron micrograph ofa 7-day sample. No poststain. Most of the strands ofpolysaccharide-like material (P) appear parallel with the artificial surface (AS). This surjace to which the microorganisms (M) are attached can be seen in the lower right corner of the electron micrograph. X 19,400.

4 VOL. 99, 1969 ELECTRON MICROSCOPIC STUDY OF SLIME LAYER 319 w ''4r.-: ^ _,!_w U1; C & *v S. lj + ' T*_v 7 t V taf-;7 i} #Se- 3-1'\ I -V 4.4. I. J% mmpw.m I m la (' - FIG. 3. View of a 9-day culture showing compacted strands of material surrounding the organisms. Note void areas surrounding most cells and the thick deposits ofpolysaccharide-like material as contrasted with Fig. I and 2. Section poststained with lead citrate. X 35,300.

5 320 JONES, ROTH, AND SANDERS J. BACTERIOL. FiG. 4. Electron micrograph of a stage of slime production, in a 9-day sample, characterized by void areas (V) immediately surrounding the cells, with closely packed strands of material surrounding the void areas. Interlaced polysaccharide-like material (P) is found between cells. Poststained with lead citrate. Marker represents I pm. X 43,000. f.o A - a...w At. _ I *,... k. ~~~~~~ I..k..It.Jr~~~~~~~~~~~~~a.O.5pM. FIG. 5. High magnification of the second general stage of slime production found in a 9-day sample. Polysaccharide-like material (P) seems to be streaming from the cell wall. Note cell membrane (CM) and absence of void area evident in Fig. 3 and 4. Poststained with lead citrate and uranyl acetate. X 105,000.

6 VOL ELECTRON MICROSCOPIC STUDY OF SLIME LAYER 321 FIG. 6. Electron micrograph of the same general stage of slime production, in a 9-day sample, as shown in Fig. 5. Note the absence of a void area. Poststained with lead citrate. X 48,000. CM 0.5)JM FiG. 7. Electron micrograph of a 9-day sample contrasting a cell with ruthenium red-stained polysaccharide layer with a cell with no apparent staining of surrounding area. The organism without the slime layer possesses a bleb (unmarked arrow) on the outer surface. Note the concentric arrangement of the strands around one organism. Poststained with lead citrate. X 67,000.

7 322 JONES, ROTH, AND SANDERS I) 2,g J. BACTERIOL. FIG. 8. Area of 9-day sample showing cells surrounded by void areas and the interlacing slime connecting each cell to give form to the slime layer. Poststained with lead citrate and uranyl acetate. X 59,000. >AS.a-; *. - ~~I b>:... P 9 M~ FIG. 9. Electron micrograph of two microorganisms in a 9-day culture which appear to be polarly oriented to the surface (AS). Note the polysaccharide-like material joining the two cells. Poststained with lead citrate and uranyl acetate. X 57,700.

8 VOL. 99) 1969 ELECTRON MICROSCOPIC STUDY OF SLIME LAYER 323 FIG. 10. High magnification of a 9-day sample showing polysaccharide-like strands attached to surface. Poststained with lead citrate and uranyl acetate. X 76,000. hope that a workable method for observing the system could be utilized to study and better understand the workings of the community. Initially, bacteria were permitted to attach to glass microscope slides submerged in the continuous-flow growth chambers for various periods of time. With this procedure, the organisms were embedded in Maraglas (17) directly on the slide after it had been removed from the chamber (6, 7, 15). After the Maraglas had polymerized, the embedded samples were pried off the slide, reembedded, thin-sectioned, and examined with an RCA EMU-2 electron microscope. Both the electron microscopic examination of the thin sections and the light microscopic examination of the glass slide after specimen removal indicated that, in some areas, the attaching layer was removed intact although there were noticeable variations in the thickness of the bacterial mass. Since it was believed that a loss of slime mass occurred during the physical manipulation of the slide before and during embedding, as well as during the removal process, the more practical embedding method reported in this paper was developed. The current view on the action of ruthenium red, as postulated by Luft (11), entails the formation of an electron-dense network with acid mucopolysaccharide when osmium tetroxide is present. Most work thus far has been done on mammalian cells surrounded by a matrix of acid mucopolysaccharide, but Pate and Ordal have used ruthenium red to stain Chondrococcus columnaris in pure culture (13). Ruthenium red appears to require the presence of OS04 in order to form an electron-dense complex with polysaccharide. Micrographs taken of sections obtained from samples stained with ruthenium red alone or with OS04 alone do not reveal the electron-dense area surrounding bacterial cells. In contrast, micrographs taken of samples in which ruthenium red was used in conjunction with OS04 show dense deposits of polysaccharide around bacterial cells. Therefore, the material stained by ruthenium red in conjunction with OS04 will be referred to in this paper as polysaccharide-like. Most cells in the 9-day samples appear to be in what is perhaps a state of low metabolic activity (those surrounded by a clear zone and then dense polysaccharide; Fig. 4), whereas a few cells seem to be actively producing strands of material and the cells stain more densely (Fig. 5 and 6). Those cells surrounded by compacted polysaccharidelike material do not stain heavily, perhaps indicating a problem with diffusion of stain through the surrounding material. These micrographs are typical of the two general stages of slime produc-

9 324 JONES, ROTH, AND SANDERS J. BACTERIOL. tion of the microorganisms found throughout the slime layer. Spaces are evident between cells, demonstrating that the slime layer of the 9-day samples is not composed of a gelatinous mat of uniform height, but rather that each cell is autonomous within its own matrix and this matrix usually is joined with a neighboring cell matrix. The horizontal view of the 9-day sample sections is generally one of an undulating mass of cells and surrounding material creating a weblike structure. Perhaps the cells surrounded by clear, nonstaining areas (Fig. 4, 7, and 8) have completed slime production. This zone may represent a microenvironment surrounding each cell, with the polysaccharide strands acting as a gradient through which diffusible and readily usable nutrients pass to the void area surrounding each cell. This microenvironment could serve as a buffer zone protecting the cell from physical damage. An artifact of dehydration may have contributed to the appearance of the void area around the cells which seem to have completed slime production. If this were the case, however, we could expect compacted slime around all organisms, including those producing slime (Fig. 5 and 6). Thus, we do not consider the void area an artifact of our specimen preparation procedure. Future samples embedded in water-soluble resin may clarify this matter (8). This void area has been noted in sections of Zoogloea made from an embedded pure culture (4). Figures 5 and 6 differ from Fig. 4 and 8 in regard to the distribution of the slime strands surrounding the microorganism. The slime may be compared to a bacterial colony in which all cells are not the same age and thus are all not in the same physiological state. The cells in Fig. 5 and 6 appear to be in a state of active polysaccharide production, whereas the cells in Fig. 3, 4, and 7 through 10 may have completed polysaccharide production and a compacting of the polysaccharide has taken place. The concentric layering seen in Fig. 7 (P) seems to indicate that in this case the polysaccharide compacted in the layer originated from this organism. The concentric layering would be difficult to explain if the polysaccharide had originated from other organisms. Generally, the compacted polysaccharide does not show such layering (Fig. 3, 4, and 8). The mechanism of this compacting is not known. Perhaps changes in ph as the cells age bring about further polymerization of the polysaccharides. Micrographs were taken of the area of attachment of microorganisms with the surface in search of specialized attachment structures. Figures 9 and 10 are representative of organisms in close apposition to the surface, and they show no specialized structures adapted for the purpose of attachment. These microorganisms seem to remain attached to the surface via a polysaccharidelike matrix in the same manner as they are attached to each other. Serial sections of these organisms do not reveal any specialized attachment structures other than the surrounding matrix. Although we have not made an extensive survey of the properties of plastics or epoxy resins which might selectively permit only certain organisms to colonize the disc surface, we have used Maraglas 655, Maraglas 732, and Epon 812 and have noted no marked difference between the quantity or quality of slime adhering to each. We are now involved in a study of the numbers and types of organisms which colonize Epon 812. Eventually, the other epoxies will be investigated in the same manner. We have made no attempt to evaluate the effect of charges on the disc surface on the selection of organisms in the slime mass. The method described in this paper is currently being used to investigate various aspects of attached organisms in streams. A suitable platform has been devised on which a slide has been mounted in a stream current, making it possible to sample directly from nature and to investigate changes in structure of indigenous organisms as opposed to growth of organisms under laboratory conditions. Another application of this method may be in the study of pure cultures under natural nutrient conditions; this can be accomplished by inserting slides in dialysis tubing containing pure cultures and placing these in a stream (5). The micrographs demonstrate that microorganisms can be studied to high levels of magnification with this method. Embedding techniques commonly employed can be readily adapted to this method, and, although sectioning occurs through two layers of epoxy resin with varying densities, suitable sections can be obtained for observation in the electron microscope. It should be noted that electron microscopic studies with ruthenium red point the way toward possible study of microbial capsule formation and structure. LITERATURE CITED 1. Cooke, W. B Colonization of artificial bare areas by microorganisms. Bot. Rev. 22: Erlanson, R. A A new maraglas D.E.R. 732 embedment for electron microscopy. J. Cell Biol. 22: Freeman, J., and B. Spurlock A new epoxy embedment for electron microscopy. J. Cell Biol. 13: Friedman, B. A., P. R. Dugan, R. M. Pfister, and C. C. Remsen Fine structure and composition of the zooloeal matrix surrounding Zoogloea ramigera. J. Bacteriol. 96: Hendricks, C. W., and S. M. Morrison Multiplication and growth of selected enteric bacteria in clear mountain stream water. Water Res. 1:

10 VOL. 99, 1969 ELECIRON MICROSCOPIC STUDY OF SLIME LAYER Heyner, S In situ embedding of cultured cells or tissue, Itrown on glass, in epoxy resins for electron microscopy. Stain Technol. 38: Howatson, A. F., and J. D. Almeida A method for the study of cultured cells by thin sectioning and electron microscopy. J. Biophys. Biochem. Cytol. 4: Kay, D. H. (ed.) Techniques for electron microscopy, 2nd ed., p F. A. Davis Co., Philadelphia. 9. Kellenberger, E., A. Ryter, and J. Sechaud Electron microscope study of DNA-containing plasms. II. Vegetative and mature phage DNA as compared with normal bacterial nucleoids in different physiological states. J. Biophys. Biochem. Cytol. 4: Luft, J. H Improvements in epoxy resin embedding methods. J. Biophys. Biochem. Cytol. 9: Luft, J. H Selective staining of acid mucopolysaccharides by ruthenium red, p In C. J. Arceneaux (ed.), Proceedings of the Electron Microscopy Society of America, 26th Annual Meeting. Claitor's Publishing Division, Baton Rouge, La. 12. Mercer, E. H., and R. J. Goldacre A technique for studying the behavior of the cell membrane in the electron microscope using alcian blue as a label, p In R. Uyeda (ed.), Electron microscopy, vol. 2. Sixth International Congress for Electron Microscopy (Kyoto, Japan). Maruzen Co., Ltd., Tokyo. 13. Pate, J. L., and J. E. Ordal The fine structure of Chondrococcus columnaris. III. The surface layers of Chondrococcus columnaris. J. Cell Biol. 35: Reynolds, E The use of lead citrate at high ph as an electron-opaque stain in electron microscopy. J. Cell Biol Robbins, E., and N. K. Gonatas In vitro selection of the mitotic cell for subsequent electron microscopy. J. Cell Biol. 20: Sanders, W. M., III Oxygen utilization by slime or ganisms in continuous culture. Air Water Poliut. Int. J. 10: Spurlock, R., V. Kattine, and J. Freeman Technical modifications in maraglas embedding. J. Cell Biol. 17: Watson, M. L Staining of tissue sections for electron microscopy with heavy metals. J. Biophys. Biochem. Cytol. 4: Wuhrmann, K River bacteriology and the role of bacteria in self-purification of rivers, p In H. Heukelekian and N. C. Dondero (ed.), Principles and applications in aquatic microbiology. John Wiley & Sons, Inc., New York.