Observations on Estuarine Microfouling Using the Scanning Electron Microscope LouIs H. D1SALVO AND G. WAYNE DANIELS Naval Biomedical Research Laboratory, University of California, Berkeley, Naval Supply Center, Oakland, California 94625
Abstract Scanning electron microscopy was used to observe microbiological primary fouling of glass surfaces exposed in estuarine waters. Observations on clean glass, and glass treated with water-repellent coatings, showed that bacterial slimes adhered less strongly to the waterrepellent glass. An experiment using pure cultures of bacteria and latex particles showed that attached bacteria promoted the settlement of latex particles on the glass.
Introduction The fouling of clean surfaces immersed in seawater begins with the attachment of microscopic particles. Nonliving particles are attracted to surfaces by complex interacting forces including electrostatic attraction, Van Der Waals forces, and polymeric bridging [6]. Bacteria and other microorganisms are subject to these (net) attraction energies and, in addition, may secrete mucilagenous adhesives in their varied strategies of surface attachment [2]. Dependent on environmental conditions and type of surface, microorganisms may multiply and adhere to the surface, producing more or less firmly anchored primary slime films. Metazoan larvae may settle on the surface, and within a period of days or weeks, the surface becomes a diversified "microbial seascape" [9]. Studies at our laboratory have investigated the occurrence and control of early events in the sequence in which bacteria,diatoms, and protozoans foul optical surfaces immersed in the sea [3]. Naval research on water-repellent flurosilane coatings designed to produce low interfacial tension at the glass-water interface showed that treated glass attracted and held more bacteria and inert particles than did clean glass surfaces, per unit area [3]. Thus, water repellency was not a good criterion of the resistance of a glass surface to initial attachment of slime formers. Empirical observations did suggest, however, that actual growth and adhesion of the lsime films was retarded on the treated glass surfaces. 234 MICROBIAL ECOLOGY, Vol. 2, 234-240 9 1975 by Springer-Verlag New York Inc.
SEM of Estuarine Microfouling
235
Often we were unable to distinguish bacteria from inert particles by light microscopy
because
of particle aggregation
a n d a l t e r a t i o n s in m o r -
p h o l o g y d u e to t h e s e c r e t i o n o f e x t r a c e l l u l a r a d h e s i v e s . T h e r e s u l t s o f T o d d et at. [ 1 0 ] a n d P a e r l [ 7 ] e n c o u r a g e d t h e u s e o f t h e s c a n n i n g e l e c t r o n m i c r o s c o p e ( S E M ) f o r o u r o b s e r v a t i o n s . I n s i t u s t u d i e s , s u c h as t h a t o f G r a y [ 5 ] , s u g g e s t e d t h a t o b s e r v a t i o n o f t h e m i c r o f o u l e r s a t t a c h e d to g l a s s u n d e r natural conditions would provide more realistic data on estuarine microbial ecology than observations of particles concentrated by filtration [8]. This contention has been amply substantiated with the recent publication of a s c a n n i n g e l e c t r o n m i c r o g r a p h s u r v e y o f v a r i o u s s u r f a c e s i m m e r s e d in s e a w a t e r [ 9 ] . T h i s p a p e r p r e s e n t s s o m e r e s u l t s o b t a i n e d in o b s e r v a t i o n o f estuarine microfouling on both water-repellent treated and untreated glass s u r f a c e s u s i n g t h e S E M . W e a l s o a t t e m p t e d s o m e e x p e r i m e n t a l w o r k to demonstrate mechanisms whereby bacteria facilitate the adhesion of inert p a r t i c l e s to t h e g l a s s .
Materials and Methods Microscope slides were rinsed in warm chromic acid cleaning solution, washed copiously with tap water, given a 1 rain rinse in distilled water, and wiped dry. These slides were designated " c l e a n " (CL). "Treated" (TR) slides were CL slides coated with SC-87 (Pierce Chemical Co., Rockford, I11.), wiped of all excess coating, and polished until crystal clear. Water contact angles on slides (see Baier et al. [1]) were approximately 20~ for CL and 90 ~ for TR; the approximate critical surface tension of CL slides was 45, and for TR, 30 dynes/cm 2 (determinations by H.R. Bleile, Mare Island Naval Ship Yard, Ca., and by J. Montemarano, Naval Ship Research and Development Center, Annapolis, Md.). When dipped in water, CL slides retained a thin water film, and TR slides drained in less than 1 min, leaving an apparently dry surface. Bacterial fouling of glass slides was accomplished in the laboratory by incubation of the slides in raw seawater at 15~ for periods of 1 to 3 days. Microfouling was obtained on slides submerged in dockside waters at the Naval Supply Center pier, Oakland, and in a sewage-polluted harbor in Freeport, Bahamas. Some films grown on slides in the laboratory were produced by inoculation of the slides with a marine bacterial species isolated from fouling plates and maintained in pure culture in the laboratory (Strain F-I). After removal from dock water or laboratory culture vessel, all slides were rinsed with approximately 100 ml filtered (0.45 /zm Millipore) seawater (MFSW) and placed in 1% glutaraldehyde or 10% formalin in MFSW, allowed to fix for at least 15 min, and then transferred to MFSW for an additional 15 min. The slides were finally washed with distilled water and air-dried. The slides were cut into 1 cm2 chips and gold-coated by vacuum deposition (approx. 100 A thickness). Samples were viewed with an Autoscan scanning electron microscope (ETEC Corp., Hayward, Ca.) using an accelerating voltage of 20 KV and incidence angle of 45 ~ Photographs were taken using the microscope's camera accessory with Polaroid No.55 film. Experimental particle association trials were carried out in which CL slides were suspended in seawater containing IOn-10a viable bacterial cells/ml, followed by washing in MFSW and immersion in a 10s-10 s particles/ml suspension of polystyrene latex (0.8 p,m diam., Dow Chemical Co.). Slides were again washed with MFSW, fixed in 10% formalin for 10 min, and finally rinsed in distilled water and air-dried. A reciprocal experiment was carried out in which slides were first placed in the latex suspension and then in the bacterial
236
L.H. DiSalvo and G. W. Daniels
suspension. A quantitative estimation was carried out with the SEM in which representative fields on slides of both trials were observed at 4000x to determine relationships between attached latex and bacteria. Twenty fields on each of two replicate slides were recorded for each experiment. To qualify for counting, a field was required to contain both bacteria and latex particles. A (+) was scored for each field in which a latex particle had adhered to one or more bacterial cells. When a field contained both latex particles and bacterial cells which were not associated, a (-) was scored. Fields containing both (+) and (-) observations were disregarded. The percentage of fields where latex particles were associated with bacterial particles was calculated for each experiment.
Results and Discussion Bacteria adhered to and formed films on both treated and clean glass slides. Figure I(A) shows bacteria in a 2-day primary film cultured under laboratory conditions on clean glass, showing a typically random distribution of cells. Under the same conditions, although growth of a bacterial film occurred, it was unable to maintain attachment to the glass as demonstrated by light microscopy [Fig. I ( B ) ] . Rolling up of these cells into intertwined ridges is shown in Fig. I(C). Figure I(D) shows a bacterial lawn and colony on a clean glass slide. Impairment o f bacterial adhesion on treated glass [Fig. I ( E ) ] . was suggested by edge retraction o f the bacterial colony after d r y i n g during the p r e p a r a t i o n for m i c r o s c o p y . Critical point drying techniques which do not stress the preparation would probably not have allowed us to discover this phenomenon. The retraction p h e n o m e n o n was routinely visible on dozens o f bacteria-fouled treated slides and never seen on clean slides. Study o f particle attachment to bacteria using pure cultures and latex showed that when slides were first immersed in bacteria, followed by immersion in latex particles, fields containing both bacteria and latex showed 95% incidence o f latex attached to bacterial cells [Fig. 2 ( A ) ] . In reciprocal trials where slides were first immersed in latex followed by immersion in bacteria, latex was associated with bacteria in only 40% of the cases observed (40 fields) [Fig. 2(B)]. Since latex particles are negatively charged, as are bacterial ceils, electrostatic attraction between these particles was ruled out. A likely explanation o f the attraction between these particles is polymer bridging, as discussed by Harris and Mitchell [ 6 ] . Following initial attraction o f latex particles to the bacteria, the particles appeared to be firmly embedded in bacterial extracellular secretions [Fig. 2(C)]. Extrapolating these results to microfouling in the field, it is apparent that inert particles may readily settle on, and be firmly attached to, bacterial colonies developing on substrates, resulting in the formation of detrital clumps as illustrated by Sieburth [ 9 ] . The phenomenon o f colony retraction was used in tentative identification o f microfouling seen on repellent-treated slides exposed in the fouling
SEM of Estuarine Microfouling
237
Fig, 1. (A) Estuarine bacteria cultured on a clean glass surface for 48 hr in a laboratory aquarium containing San Francisco Bay water (from 4000• (B) Bacterial film on water repellent-treated glass surface after 48 hr in laboratory aquarium. Semicoherent film is seen to left of center (f), areas showing nonadhesion of the bacterial film (c) contain a few bacterial single cells (from 4000• (C) Estuarine bacteria on water repellent-treated glass from B (above). Cells fail to spread on glass, as do extracellular adhesives (arrows) (from 4000x). (D) A colony and " l a w n " of an estuarine bacterial isolate grown on a clean glass slide in the laboratory (from 4000• (E) Colony edge of same isolate as D (above) grown on water repellent-treated glass, showing typical retraction zone (arrow) (from 4000•
238
L.H. DiSalvo and G. W. Daniels
study carried out in the Bahamas in February, 1974. Figure 3(A) shows a clump of particles adherent to a TR slide which had been immersed for 6 days in a sewage-polluted harbor. Under light microscopy this clump would have been recorded as "particulate organic detritus." Careful observation at 4000• with the SEM showed a retracted edge [3(B)] as described above for pure culture colonies on TR slides. The clump was thus presumeably a bacterial colony covered with attached particles. A primary problem in the use of the SEM in observation of samples obtained in nature was the inability to identify microfouling organisms growing on the slides. Figures 4(A) and 4(B) illustrate two as yet unidentified organisms commonly found on the fouling slides. It is envisioned that significant numbers of the organisms seen in the microfouling sequence will be identified once they have been isolated from microcolonies, grown under
Fig. 2. (A) Clean glass surface which had been first immersed in a bacterial suspension followed by rinsing and immersion in a suspension of polystyrene latex particles (1 , latex; b , bacterial cells) (from 4000x). (B) Same as above, with slide first immersed in latex particle suspension, followed by rinse, and immersion in bacterial suspension (a , crystalline artifacts encountered in drying process) (from 4000• (C) Latex particles (I) attached to bacterial aggregation (b) (from 8000•
SEM of Estuarine Microfouling
239
pure culture c o n d i t i o n s , and c h a r a c t e r i z e d by light m i c r o s c o p y and scanning e l e c t r o n m i c r o s c o p y . H o w e v e r , o r g a n i s m s w h i c h are c u l t u r e d in the l a b o r a t o r y m a y not n e c e s s a r i l y exhibit the s a m e m o r p h o l o g y as those grown under field c o n d i t i o n s , and it is indeed l i k e l y that s o m e o f the bizarre forms may not be a m e n a b l e to l a b o r a t o r y culture.
Fig. 3. (A) Aggregate of particulate detritus attached to a water repellent-treated glass slide immersed in a polluted harbor for 6 days near Lucaya, Bahamas, in Feb., 1974. Area at (1) is magnified in Fig. 3 (B), and organism at (2) in Fig. 4 (A) (from 2000x). (B) Margin of detrital aggregate (A) demonstrating retraction zone commonly seen for bacterial colonies grown in pure culture on treated surfaces [Fig. I (E)] [d , diatom frustule; b , bacterial single cell; c , bacterial colony cell (Tentative ID)J (from 8000 •
Fig. 4. (A) Unidentified microorganism (center) attached to fouling slide immersed in a polluted harbor in the Bahamas for 6 days in Feb., 1974. Bacterial cells at arrows; d , diatom frustules (from 8000• (B) Unidentified microorganism colony developing on a glass surface immersed in San Francisco Bay for 5 days in Fall, 1973 (from 4000•
240
L.H.
DiSalvo and G. W. Daniels
Acknowledgments We gratefully acknowledge support of this work by the Office of Naval Research and the Manned Undersea Science Project of the National Oceanic and Atmospheric Administration. I thank my colleague Dr. H. C. Ross and his wife Glenda for the valuable support rendered in the Bahamas.
References 1.
Baier, R. E., Shafrin, E. G., and Zisman, W. A. 1968. Adhesion; Mechanisms that assist or impede it. Science 162: 1360.
2.
Corpe, W. A. 1970. Attachment of marine bacteria to solid surfaces. In: Adhesion in Biological Systems. R. Manly, editor Pp. 7 3 - 8 5 . Academic Press, New York.
3.
Di Salvo, L. H. 1972. Early steps in the microbial fouling of optical surfaces in the marine environment. In: 47th Ann. Tech. Prog. Rept., Naval Biomed. Res. Lab. U. C. Berkeley. Pp. 357-378.
4.
Di Salvo, L. H., and Cobet, A. B. 1974. Control of an estuarine microfouling sequence on optical surfaces using low intensity ultraviolet irradiation. Appl. Microbiol. 27: 172.
5.
Gray, T. R. G. 1967. Stereoscan electron microscopy of soil microorganisms. Science 155: 1668.
6.
Harris, R. H., and Mitchell, R. 1973. The role of polymers in microbial aggregation. In: Ann. Rev. Microbiol. M. P. Start et al., editors. Annual Reviews Inc. Pp. 27-50. Palo Alto, California.
7.
Paerl, H. W. 1973. Detritus in Lake Tahoe: Structural modification by attached microflora. Science 180: 496.
8.
Paerl, H. W., and Shimp, S. L. 1973. Preparation of filtered plankton and detritus for study with scanning electron microscopy. Limnol. Oceanogr. 18: 802.
9.
Sieburth, J. M. 1975. Microbial Seascapes. University Park Press, Baltimore.
10.
Todd, R. L., Humphreys, W. J., and Odum, E. P. 1973. The application of scanning electron microscopy to estuarine microbial research. In: Belle Baruch Symposium. Vol. 1, Pp. 115-125. L. H. Stevenson and R. R. Colwell, editors. South Carolina Press, Columbia.
Abstract Scanning electron microscopy was used to observe microbiological primary fouling of glass surfaces exposed in estuarine waters. Observations on clean glass, and glass treated with water-repellent coatings, showed that bacterial slimes adhered less strongly to the waterrepellent glass. An experiment using pure cultures of bacteria and latex particles showed that attached bacteria promoted the settlement of latex particles on the glass.
Introduction The fouling of clean surfaces immersed in seawater begins with the attachment of microscopic particles. Nonliving particles are attracted to surfaces by complex interacting forces including electrostatic attraction, Van Der Waals forces, and polymeric bridging [6]. Bacteria and other microorganisms are subject to these (net) attraction energies and, in addition, may secrete mucilagenous adhesives in their varied strategies of surface attachment [2]. Dependent on environmental conditions and type of surface, microorganisms may multiply and adhere to the surface, producing more or less firmly anchored primary slime films. Metazoan larvae may settle on the surface, and within a period of days or weeks, the surface becomes a diversified "microbial seascape" [9]. Studies at our laboratory have investigated the occurrence and control of early events in the sequence in which bacteria,diatoms, and protozoans foul optical surfaces immersed in the sea [3]. Naval research on water-repellent flurosilane coatings designed to produce low interfacial tension at the glass-water interface showed that treated glass attracted and held more bacteria and inert particles than did clean glass surfaces, per unit area [3]. Thus, water repellency was not a good criterion of the resistance of a glass surface to initial attachment of slime formers. Empirical observations did suggest, however, that actual growth and adhesion of the lsime films was retarded on the treated glass surfaces. 234 MICROBIAL ECOLOGY, Vol. 2, 234-240 9 1975 by Springer-Verlag New York Inc.
SEM of Estuarine Microfouling
235
Often we were unable to distinguish bacteria from inert particles by light microscopy
because
of particle aggregation
a n d a l t e r a t i o n s in m o r -
p h o l o g y d u e to t h e s e c r e t i o n o f e x t r a c e l l u l a r a d h e s i v e s . T h e r e s u l t s o f T o d d et at. [ 1 0 ] a n d P a e r l [ 7 ] e n c o u r a g e d t h e u s e o f t h e s c a n n i n g e l e c t r o n m i c r o s c o p e ( S E M ) f o r o u r o b s e r v a t i o n s . I n s i t u s t u d i e s , s u c h as t h a t o f G r a y [ 5 ] , s u g g e s t e d t h a t o b s e r v a t i o n o f t h e m i c r o f o u l e r s a t t a c h e d to g l a s s u n d e r natural conditions would provide more realistic data on estuarine microbial ecology than observations of particles concentrated by filtration [8]. This contention has been amply substantiated with the recent publication of a s c a n n i n g e l e c t r o n m i c r o g r a p h s u r v e y o f v a r i o u s s u r f a c e s i m m e r s e d in s e a w a t e r [ 9 ] . T h i s p a p e r p r e s e n t s s o m e r e s u l t s o b t a i n e d in o b s e r v a t i o n o f estuarine microfouling on both water-repellent treated and untreated glass s u r f a c e s u s i n g t h e S E M . W e a l s o a t t e m p t e d s o m e e x p e r i m e n t a l w o r k to demonstrate mechanisms whereby bacteria facilitate the adhesion of inert p a r t i c l e s to t h e g l a s s .
Materials and Methods Microscope slides were rinsed in warm chromic acid cleaning solution, washed copiously with tap water, given a 1 rain rinse in distilled water, and wiped dry. These slides were designated " c l e a n " (CL). "Treated" (TR) slides were CL slides coated with SC-87 (Pierce Chemical Co., Rockford, I11.), wiped of all excess coating, and polished until crystal clear. Water contact angles on slides (see Baier et al. [1]) were approximately 20~ for CL and 90 ~ for TR; the approximate critical surface tension of CL slides was 45, and for TR, 30 dynes/cm 2 (determinations by H.R. Bleile, Mare Island Naval Ship Yard, Ca., and by J. Montemarano, Naval Ship Research and Development Center, Annapolis, Md.). When dipped in water, CL slides retained a thin water film, and TR slides drained in less than 1 min, leaving an apparently dry surface. Bacterial fouling of glass slides was accomplished in the laboratory by incubation of the slides in raw seawater at 15~ for periods of 1 to 3 days. Microfouling was obtained on slides submerged in dockside waters at the Naval Supply Center pier, Oakland, and in a sewage-polluted harbor in Freeport, Bahamas. Some films grown on slides in the laboratory were produced by inoculation of the slides with a marine bacterial species isolated from fouling plates and maintained in pure culture in the laboratory (Strain F-I). After removal from dock water or laboratory culture vessel, all slides were rinsed with approximately 100 ml filtered (0.45 /zm Millipore) seawater (MFSW) and placed in 1% glutaraldehyde or 10% formalin in MFSW, allowed to fix for at least 15 min, and then transferred to MFSW for an additional 15 min. The slides were finally washed with distilled water and air-dried. The slides were cut into 1 cm2 chips and gold-coated by vacuum deposition (approx. 100 A thickness). Samples were viewed with an Autoscan scanning electron microscope (ETEC Corp., Hayward, Ca.) using an accelerating voltage of 20 KV and incidence angle of 45 ~ Photographs were taken using the microscope's camera accessory with Polaroid No.55 film. Experimental particle association trials were carried out in which CL slides were suspended in seawater containing IOn-10a viable bacterial cells/ml, followed by washing in MFSW and immersion in a 10s-10 s particles/ml suspension of polystyrene latex (0.8 p,m diam., Dow Chemical Co.). Slides were again washed with MFSW, fixed in 10% formalin for 10 min, and finally rinsed in distilled water and air-dried. A reciprocal experiment was carried out in which slides were first placed in the latex suspension and then in the bacterial
236
L.H. DiSalvo and G. W. Daniels
suspension. A quantitative estimation was carried out with the SEM in which representative fields on slides of both trials were observed at 4000x to determine relationships between attached latex and bacteria. Twenty fields on each of two replicate slides were recorded for each experiment. To qualify for counting, a field was required to contain both bacteria and latex particles. A (+) was scored for each field in which a latex particle had adhered to one or more bacterial cells. When a field contained both latex particles and bacterial cells which were not associated, a (-) was scored. Fields containing both (+) and (-) observations were disregarded. The percentage of fields where latex particles were associated with bacterial particles was calculated for each experiment.
Results and Discussion Bacteria adhered to and formed films on both treated and clean glass slides. Figure I(A) shows bacteria in a 2-day primary film cultured under laboratory conditions on clean glass, showing a typically random distribution of cells. Under the same conditions, although growth of a bacterial film occurred, it was unable to maintain attachment to the glass as demonstrated by light microscopy [Fig. I ( B ) ] . Rolling up of these cells into intertwined ridges is shown in Fig. I(C). Figure I(D) shows a bacterial lawn and colony on a clean glass slide. Impairment o f bacterial adhesion on treated glass [Fig. I ( E ) ] . was suggested by edge retraction o f the bacterial colony after d r y i n g during the p r e p a r a t i o n for m i c r o s c o p y . Critical point drying techniques which do not stress the preparation would probably not have allowed us to discover this phenomenon. The retraction p h e n o m e n o n was routinely visible on dozens o f bacteria-fouled treated slides and never seen on clean slides. Study o f particle attachment to bacteria using pure cultures and latex showed that when slides were first immersed in bacteria, followed by immersion in latex particles, fields containing both bacteria and latex showed 95% incidence o f latex attached to bacterial cells [Fig. 2 ( A ) ] . In reciprocal trials where slides were first immersed in latex followed by immersion in bacteria, latex was associated with bacteria in only 40% of the cases observed (40 fields) [Fig. 2(B)]. Since latex particles are negatively charged, as are bacterial ceils, electrostatic attraction between these particles was ruled out. A likely explanation o f the attraction between these particles is polymer bridging, as discussed by Harris and Mitchell [ 6 ] . Following initial attraction o f latex particles to the bacteria, the particles appeared to be firmly embedded in bacterial extracellular secretions [Fig. 2(C)]. Extrapolating these results to microfouling in the field, it is apparent that inert particles may readily settle on, and be firmly attached to, bacterial colonies developing on substrates, resulting in the formation of detrital clumps as illustrated by Sieburth [ 9 ] . The phenomenon o f colony retraction was used in tentative identification o f microfouling seen on repellent-treated slides exposed in the fouling
SEM of Estuarine Microfouling
237
Fig, 1. (A) Estuarine bacteria cultured on a clean glass surface for 48 hr in a laboratory aquarium containing San Francisco Bay water (from 4000• (B) Bacterial film on water repellent-treated glass surface after 48 hr in laboratory aquarium. Semicoherent film is seen to left of center (f), areas showing nonadhesion of the bacterial film (c) contain a few bacterial single cells (from 4000• (C) Estuarine bacteria on water repellent-treated glass from B (above). Cells fail to spread on glass, as do extracellular adhesives (arrows) (from 4000x). (D) A colony and " l a w n " of an estuarine bacterial isolate grown on a clean glass slide in the laboratory (from 4000• (E) Colony edge of same isolate as D (above) grown on water repellent-treated glass, showing typical retraction zone (arrow) (from 4000•
238
L.H. DiSalvo and G. W. Daniels
study carried out in the Bahamas in February, 1974. Figure 3(A) shows a clump of particles adherent to a TR slide which had been immersed for 6 days in a sewage-polluted harbor. Under light microscopy this clump would have been recorded as "particulate organic detritus." Careful observation at 4000• with the SEM showed a retracted edge [3(B)] as described above for pure culture colonies on TR slides. The clump was thus presumeably a bacterial colony covered with attached particles. A primary problem in the use of the SEM in observation of samples obtained in nature was the inability to identify microfouling organisms growing on the slides. Figures 4(A) and 4(B) illustrate two as yet unidentified organisms commonly found on the fouling slides. It is envisioned that significant numbers of the organisms seen in the microfouling sequence will be identified once they have been isolated from microcolonies, grown under
Fig. 2. (A) Clean glass surface which had been first immersed in a bacterial suspension followed by rinsing and immersion in a suspension of polystyrene latex particles (1 , latex; b , bacterial cells) (from 4000x). (B) Same as above, with slide first immersed in latex particle suspension, followed by rinse, and immersion in bacterial suspension (a , crystalline artifacts encountered in drying process) (from 4000• (C) Latex particles (I) attached to bacterial aggregation (b) (from 8000•
SEM of Estuarine Microfouling
239
pure culture c o n d i t i o n s , and c h a r a c t e r i z e d by light m i c r o s c o p y and scanning e l e c t r o n m i c r o s c o p y . H o w e v e r , o r g a n i s m s w h i c h are c u l t u r e d in the l a b o r a t o r y m a y not n e c e s s a r i l y exhibit the s a m e m o r p h o l o g y as those grown under field c o n d i t i o n s , and it is indeed l i k e l y that s o m e o f the bizarre forms may not be a m e n a b l e to l a b o r a t o r y culture.
Fig. 3. (A) Aggregate of particulate detritus attached to a water repellent-treated glass slide immersed in a polluted harbor for 6 days near Lucaya, Bahamas, in Feb., 1974. Area at (1) is magnified in Fig. 3 (B), and organism at (2) in Fig. 4 (A) (from 2000x). (B) Margin of detrital aggregate (A) demonstrating retraction zone commonly seen for bacterial colonies grown in pure culture on treated surfaces [Fig. I (E)] [d , diatom frustule; b , bacterial single cell; c , bacterial colony cell (Tentative ID)J (from 8000 •
Fig. 4. (A) Unidentified microorganism (center) attached to fouling slide immersed in a polluted harbor in the Bahamas for 6 days in Feb., 1974. Bacterial cells at arrows; d , diatom frustules (from 8000• (B) Unidentified microorganism colony developing on a glass surface immersed in San Francisco Bay for 5 days in Fall, 1973 (from 4000•
240
L.H.
DiSalvo and G. W. Daniels
Acknowledgments We gratefully acknowledge support of this work by the Office of Naval Research and the Manned Undersea Science Project of the National Oceanic and Atmospheric Administration. I thank my colleague Dr. H. C. Ross and his wife Glenda for the valuable support rendered in the Bahamas.
References 1.
Baier, R. E., Shafrin, E. G., and Zisman, W. A. 1968. Adhesion; Mechanisms that assist or impede it. Science 162: 1360.
2.
Corpe, W. A. 1970. Attachment of marine bacteria to solid surfaces. In: Adhesion in Biological Systems. R. Manly, editor Pp. 7 3 - 8 5 . Academic Press, New York.
3.
Di Salvo, L. H. 1972. Early steps in the microbial fouling of optical surfaces in the marine environment. In: 47th Ann. Tech. Prog. Rept., Naval Biomed. Res. Lab. U. C. Berkeley. Pp. 357-378.
4.
Di Salvo, L. H., and Cobet, A. B. 1974. Control of an estuarine microfouling sequence on optical surfaces using low intensity ultraviolet irradiation. Appl. Microbiol. 27: 172.
5.
Gray, T. R. G. 1967. Stereoscan electron microscopy of soil microorganisms. Science 155: 1668.
6.
Harris, R. H., and Mitchell, R. 1973. The role of polymers in microbial aggregation. In: Ann. Rev. Microbiol. M. P. Start et al., editors. Annual Reviews Inc. Pp. 27-50. Palo Alto, California.
7.
Paerl, H. W. 1973. Detritus in Lake Tahoe: Structural modification by attached microflora. Science 180: 496.
8.
Paerl, H. W., and Shimp, S. L. 1973. Preparation of filtered plankton and detritus for study with scanning electron microscopy. Limnol. Oceanogr. 18: 802.
9.
Sieburth, J. M. 1975. Microbial Seascapes. University Park Press, Baltimore.
10.
Todd, R. L., Humphreys, W. J., and Odum, E. P. 1973. The application of scanning electron microscopy to estuarine microbial research. In: Belle Baruch Symposium. Vol. 1, Pp. 115-125. L. H. Stevenson and R. R. Colwell, editors. South Carolina Press, Columbia.
