Effects of antibiotics on the concentration of bacteria in biofilms and on the growth of Haliotis rufescens postlarvae.
KEY WORDS: Haliotis rufescens, abalone postlarvae, antibiotics, bacteria
Intensive rearing conditions allow the increase of bacteria caused by high levels of organic matter in culture systems. In many aquaculture facilities, bacteria are controlled using different antibiotics. (Li et al. 1999, Rombaud et al. 1999, Torkildsen et al. 2000). The triumph of antibiotics over disease-causing bacteria is one of modern medicine's greatest success stories. However over time, some bacteria have developed adaptations to minimize the effects of antibiotics (Fernandez et al. 2000, Holstrom et al. 2003). The routine use of antibiotics in aquaculture may cause development of antibiotic resistance among pathogens infecting cultured animals and humans. Antibiotics may also cause negative effects on important ecosystem bacteria (Benbrook 2002, Holstrom et al. 2003, Costanzo et al. 2005).
Abalone (Haliotis spp.) postlarvae are cultured in tanks with surfaces covered with biofilms dominated by benthic diatoms that are grazed by postlarvae. Pathogenic bacteria in these biofilms can reach high concentrations and cause abalone mortality (Anguiano-Beltran et al. 1998, Lizarraga-Partida et al. 1998).
Literature on the effects of antibiotics on bacteria associated with biofilms used in the culture of abalone is lacking, although in abalone postlarval research some antibiotics are used to control or prevent bacterial problems, including a mixture of penicillin G sodium/streptomycin sulphate (50-100 mg[L.sup.-1]) and chloramphenicol (10-20 mg [L.sup.-1]), (Searcy-Bernal et al. 1992, Roberts & Nicholson 1997, Roberts et al. 2001, Searcy-Bernal et al. 2001).
The main objective of this work is to study the effects of chloramphenicol and a mixture of penicillin/streptomycin on the bacterial concentrations in biofilms of the diatom Navicula incerta, which is widely used in abalone culture. Bacteria are also considered an important food source for early abalone postlarvae (Kawamura et al. 1998) and if antibiotics decreased their concentration substantially, an effect on early postlarval growth would be expected.
MATERIALS AND METHODS
Two assays were performed to evaluate the effects of different concentrations of the antibiotics: chloramphenicol (experiment 1) and a mixture of penicillin G sodium/dihydrostreptomycin (experiment 2), on bacterial counts in biofilms associated with Haliotis rufescens culture and on the growth of postlarvae.
The Microalgae Laboratory of the Instituto de Investigaciones Oceanoldgicas provided the diatom Navicula incerta used to form the biofilm to feed abalone postlarvae. The farm "Abulones Cultivados S. A. de C. V." (Ejido Erendira, B.C., Moxico) donated the veliger larvae of Haliotis rufescens. Larvae were reared in 1-[micro]m filtered, UV treated seawater changed every day. Competent larvae (7-8 days old) were induced to metamorphose in rectangular plastic vessels (1 L) with gammaaminobutyric acid (GABA) at 1.5-[micro]M (Searcy-Bernal & Anguiano-Beltran 1998).
Sterile 12-well tissue culture plates (Corning, 3.8 [cm.sup.2] bottom area, and 6 mL volume) were used as experimental units (EU). Diatoms were inoculated at 250-cell [mm.sup.-2] one day before the beginning of the trials and 4-5 postlarvae (5-d old in experiment 1 and 3-d old in experiment 2) were transferred to each EU the next day. All antibiotics were tested at four concentrations with three replicates per treatment following a completely randomized design. Stock solutions of antibiotics were prepared with sterile distilled water and work solutions were prepared with sterile seawater both of them at the moment of use.
Before adding the antibiotics to the EU samples (1-mL) from the biofilm were collected with an automatic sterilized pipette from a previously standardized area, to estimate the initial number of bacteria. These samples were suspended in a test tube with 9 mL of saline solution (0.9% NaCl), and immersed in an ultrasound bath (Fisher Scientific) for 3 min to disaggregate diatoms and to detach bacteria from diatoms (Lizarraga et al. 1998, Gorrostieta-Hurtado & Searcy-Bernal 2004). Second and third dilutions of these sonicated test tubes were prepared and samples of these (100 [micro]L) were plated in triplicate onto Zobell agar media and incubated at 24[degress]C to 25[degrees]C for 5 days. Bacterial counts at 5 days after incubation were made to determine CFU [mL.sup.-1]. Experiments were conducted at 17 [+ or -] 1[degrees]C under constant fluorescent illumination (ca. 50 [micro]E s [m.sup.-2]).
For experiment 1, concentrations of chloramphenicol were 0, 5, 10, and 20 mg [L.sup.-1] and biofilm samples were taken at 1, 2, 7, 15, and 22 days to evaluate bacterial concentrations. In experiment 2, 0/0, 50/50, 100/100, and 150/150 mg [L.sup.-1] of penicillin G sodium/dihydrostreptomycin were used and biofilm samples were taken at days 1, 2, 7, 14, 21, 28, and 35. Antibiotics were replaced every other day during the water changes.
To measure shell length, video-images of 2-4 postlarvae from each EU were recorded with a Sony SSC-C374 high-resolution camera on an inverted microscope (Meiji Techno) and analyzed digitally (NIH image 1.59 Power PC). Survival was evaluated counting live and dead postlarvae under an inverted microscope only in the experiment 2. The statistical significance of the differences between antibiotic concentrations was determined by one-way analyses of variance. Percent survival data were subjected to the arcsine square-root transformation before the analyses (Zar 1999).
The bacterial growth in the biofilm of Navicula incerta associated with abalone postlarvae under different chloramphenicol concentrations after 22 days of experimentation is presented in Figure 1. Only at the highest antibiotic concentration, bacterial concentrations decreased to 2.2 x [10.sup.3] CFU [mL.sup.-1] in the first 24 h of treatment, but increased in all treatments the next day reaching concentrations of [10.sup.6] at the end of the experiment. Differences among treatments were significant at day 1 (F = 4.12, P = 0.05) and day 11 (F = 5.01, P = 0.03).
[FIGURE 1 OMITTED]
There was not a clear relationship between postlarval growth and chloramphenicol concentration (Fig. 2). At the end of the experiment the largest average shell length ([+ or -] s. error.) was 557.3 [+ or -] 19.1 [micro]m at 20 mg [L.sup.-1] and the smallest (472.8 [+ or -] 18.8 [micro]m) at 5 mg [L.sup.-1] and this difference was significant (F = 5.70, P = 0.02).
Bacterial counts in the experiment with the mixture of penicillin G sodium/dihydrostreptomycin are presented in Figure 3. The highest antibiotic concentration (150/150 mg [L.sup.-1]) depleted the bacterial population to 0.2 [+ or -] 0.1 x [10.sup.3] CFU [mL.sup.-1] after 48 h of treatment although differences among treatments were not significant (F = 1.0, P = 0.5). During the next three weeks there were important differences (of ca. one order of magnitude) between the control and the highest concentration, which were significant at day 7 (F = 8.05, P = 0.007) and day 14 (F = 4.4, P = 0.04). At the end of the experiment (day 35) biofilms were severely grazed by postlarvae and, probably as a consequence, bacterial concentrations decreased in all treatments (Fig. 3). The highest value (2 x [10.sup.5] CFU [mL.sup.-1]) was still in the control and the lowest in the 100/100 mg [L.sup.-1] mixture (6 x [10.sup.3] CFU [mL.sup.-1]). This difference was significant (F = 4.2, P = 0.05).
During the first three weeks, postlarval growth in the highest concentration of the antibiotic mixture (150/150 mg [L.sup.1]) was slower than in the other treatments (Fig. 4), and this difference was significant at day 21 (F = 5.4, P = 0.03). At the end of the experiment, postlarval survival was also lower (31%) in the highest antibiotic concentration (Fig. 5), and a significant difference among treatments was detected (F = 7.77, P = 0.009).
Bacteria are some of the most important agents that cause mass mortalities of animals. The use of antibiotics to control diseases in aquaculture promotes the development of resistant strains of both harmful and harmless bacteria that are replacing antibiotic susceptible bacteria (Son et al. 1997, Li et al. 1999). Fitt et al. (1992) mention that some antibiotics can control bacterial growth if added daily to culture; but in this study chloramphenicol and penicillin/dihydrostreptomycin were effective only during the first 48 h, despite continuous replacement with fresh antibiotics. These findings suggest the development of antibiotic resistance by bacteria because of the frequent use of antibiotics, in agreement with other reports (Son et al. 1997, Bruun et al. 2000, Chelossi et al. 2003). Bacteria were probably sensitive to antibiotics during the first two days, because diatoms and postlarvae never received an antibiotic treatment before.
[FIGURE 2 OMITTED]
[FIGURE 3 OMITTED]
On other hand, because diatoms and bacteria are forming a biofilm, it is possible that its complex structure, based on exopolysaccharides, provides protection to bacterial populations against antibiotics as suggested by Nickel et al. (1985), Nichols et al. (1988), and Davey and Otoole (2000).
Abalones require gut bacteria to digest algae (Erasmus et al. 1997, Sawabe et al. 1998). The digestive system of the early postlarvae is not yet well developed, thus they need to acquire the bacterial flora from the environment to feed efficiently on microalgae (Tanaka et al. 2003). Erasmus et al. (1997) found that chloramphenicol and cefotaxime at 20 and 150-[micro]g [mL.sup.-1] respectively inhibited the growth of abalone enteric bacteria.
[FIGURE 4 OMITTED]
[FIGURE 5 OMITTED]
Therefore, since in our study it was found that high levels of chloramphenicol and penicillin G sodium/dihydrostreptomycin decreased bacterial numbers during the first days of experimentation, it is possible that in these treatments postlarvae did not develop a normal gut microflora, either because of a low availability in the biofilm or the potential inhibition of bacterial growth in the digestive system. In the case of chloramphenicol, this was probably not an important issue, because the concentrations of bacteria increased quickly to control levels ([10.sup.6] CFU [mL.sup.-1]) and postlarval growth was similar to that in the control (Figs. 1, 2). However, in the highest penicillin/streptomycin (150/150 mg [L.sup.-1]) treatment, bacteria remained lower than in the control during all the experimental period and postlarval growth was also slower during the first 3-4 wk (Figs. 3, 4). The lowest survival in this high antibiotic treatment (Fig. 5) might also be related to a digestive deficiency caused by the lack of appropriate microflora. On the other hand, these results might be also partially explained by the potential role of bacteria as a nutritional source for abalone postlarvae (Kawamura et al. 1998). The potential toxic effect of high doses of antibiotics on abalone postlarvae cannot be ruled out, although literature on this issue is lacking.
In abalone research, antibiotics have been used in an effort to control bacterial interference during the chemical induction of larval metamorphosis, as well as mortality during postlarval development. These include the mixture of penicillin/streptomycin at 150/150 mg [L.sup.-1] each (Searcy-Bernal et al. 1992, Roberts et al. 2001, Takami et al. 2002) and chloramphenicol at 10 mg [L.sup.-1] (Martinez-Ponce & Searcy-Bernal 1998, Searcy-Bernal et al. 2001). Results from this study suggest that these antibiotics may be helpful in short-term studies such as those dealing with metamorphosis induction of abalone, because this process takes only a few days (Roberts & Nicholson 1997, Searcy-Bernal & Anguiano-Beltran 1998, Roberts 2001). However, this may not be the case in long-term postlarval studies, because bacterial concentrations recover in a few days, unless antibiotics affect pathogenic bacteria more efficiently and these are reduced in the final microbial composition. Gapasin & Polohan (2004) used streptomycin sulfate at 10 mg [L.sup.1] in their metamorphosis induction experiments, but it is doubtful whether this low concentration had any effect in the control of potential bacterial infection and larval mortality as intended.
This study suggests that the long-term prophylactic use of antibiotics in abalone culture systems may not be the best health practice and it is necessary to find some other alternatives, such as the use of probiotics (Douillet & Langdon 1993, Avendano & Riquelme 1999, Makridis et al. 2000, Bachere 2003). Further research is required on the effect of antibiotics on the species composition in the microbial community of biofilms.
The authors thank the farm Abulones Cultivados for the donation of abalone larvae and the Microalgae Laboratory of the I.I.O for providing the diatom Navicula incerta. They also thank two anonymous reviewers for their comments on the manuscript. This study was partially funded by the University of Baja California (grants 529 and 4403) and the Mexican Government (CONACyT grant 37461-B and SNI scholarship 5532). This paper is part of the doctoral dissertation of C. Anguiano-Beltran supported by a CONACyT scholarship.
Anguiano-Beltran, C., R. Searcy-Bernal & M. L. Lizarraga-Partida. 1998. Pathogenic effects of Vibrio alginolyticus on larvae and postlarvae of the red abalone (Haliotis rufescens). Dis. Aquat. Org. 33:111-118.
Avendano, R. E. & C. E. Riquelme. 1999. Establishment of mixed-culture probiotics and microalgae as food for viable larvae. Aquacult. Res. 30:893-900.
Bachere, E. 2003. Anti-infectious immune effectors in marine invertebrates: potential tools for disease control in larviculture. Aquaculture 227:427-438.
Benbrook, C. M. 2002. Antibiotic drug use in United States aquaculture. 2002. IATP Report. The Northwest Science and Environmental Policy Center. Sandpoint, Idaho. 16 pp.
Bruun, M. S., A. S. Schmidt, L. Madsen & I. Dalsgaard. 2000. Antimicrobial resistance patterns in Danish isolates of Flavobacterium psychrophilum. Aquaculture 187:210-212.
Chelossi, E., L. Vezzulli, A. Milano, M. Branzoni, M. Fabiano, G. Riccardi & I. M. Banat. 2003. Antibiotic resistance of benthic bacteria in fish farm and control sediments of western Mediterranean. Aquaculture 219:83-97.
Costanzo, S. D., J. Murby & J. Bates. 2005. Ecosystem response to antibiotics entering the aquatic environment. Mar. Pollut. Bull. 51:218-223.
Davey, M. E. & G. A. Otoole. 2000. Microbial biofilms: from ecology to molecular genetics. Microbiol. Mol. Biol. Rev. 64:847-867.
Douillet, P. & C. J. Langdon. 1993. Effect of marine bacteria on the culture of axenic oyster Crassostrea gigas (Thunberg) larvae. Biol. Bull. 184:36-51.
Erasmus, J. H., P. A. Cook & V. E. Coyne. 1997. The role of bacteria in the digestion of seaweed by the abalone H. midae. Aquaculture 155:377-386.
Fernandez, H., M. Mansilla & V. Gonzalez. 2000. Antimicrobial susceptibility of Campilobacter jejuni subsp. jejuni assessed by E-test and double dilution agar method in Southern Chile. Mem. Inst. Oswaldo Cruz 95:247-249. (Rio de Janeiro)
Fitt, W. K., G. A. Heslinga & T. C. Watson. 1992. Use of antibiotics in the mariculture of giant clams (F. Tridacnidae). Aquaculture 104: 1-10.
Gapasin, R. S. J. & B. B. Polohan. 2004. Induction of larval settlement and metamorphosis in the donkey-ear abalone. Haliotis asinina Linnaeus, by chemical cues. Hydrobiologia 519:9-17.
Gorrostieta-Hurtado, E. & R. Searcy-Bernal. 2004. Combined effects of light conditions (constant illumination or darkness) and diatom density on postlarval survival and growth abalone Haliotis rufescens. J. Shellfish Res. 23:1001-1008.
Holstrom, K., S. Graslund, A. Wahlstrom, S. Poungshompoo, B. Bengtsson & N. Kautsky. 2003. Antibiotic use in shrimp farming and implications for environmental impacts and human health. Int. J. Food Sci. 38:255-266.
Kawamura, T., R. Roberts & H. Takami. 1998. A review of feeding and growth of postlarval abalone. J. Shellfish Res. 17:615-625.
Li, J., Yie, R. W. Foo, J. M. L. Ling, H. Xu & N. Y. S. Woo. 1999. Antibiotic resistance and plasmid profiles of Vibrio isolates from cultured silver sea bream, Sparus sarba. Mar. Poll. Bull. 39:245-249.
Lizarraga-Partida, M. L., C. Anguiano-Beltran, R. Searcy-Bernal & E. Vazquez-Moreno. 1998. Bacterial water quality in abalone farms of Baja California. Y. Shellfish Res. 17:689-692.
Makridis, P., A. J. Fjellheim, J. Skjermo & O. Vadstein. 2000. Control of the bacterial flora of Brachionus plicatilis and Artemia franciscana by incubation in bacterial suspensions. Aquaculture 185:207-218.
Martinez-Ponce, D. & R. Searcy-Bernal. 1998. Grazing rates of red abalone (Haliotis rufescens) postlarva feeding on the benthic diatom Navicula incerta. J. Shellfish Res. 17:627-630.
Nickel, J. C., I. Ruseska. J. B. Wright & J. W. Costerton. 1985. Tobramicin resistance of Pseudomonas aeruginosa cells growing as a biofilm on urinary catheter material. Antimic. Agents Chemother. 27:619-624.
Nichols, W. W., S. M. Darrington, M. P. E. Slack & H. L. Walmsley. 1988. Inhibition of tobramycin diffusion by binding to alginate. Antimicrob. Agents Chemother. 32:518-523.
Roberts, R. D. & C. M. Nicholson. 1997. Variable response from abalone larvae (Haliotis iris, H. virginea) to a range of settlement cues. Moll. Res. 18:131-141.
Roberts, R. 2001. A review of settlement cues for larval abalone (Haliotis spp.). J. Shellfish Res. 20:571-586.
Roberts, R., C. Lapworth & R. J. Barker. 2001. Effect of starvation on the growth and survival of post-larval abalone (Haliotis iris). Aquaculture 200:323-338.
Rombaud, G., Ph. Dhert, J. Vandenberghe, L. Verschuere, P. Sorgeloos & W. Verstraete. 1999. Selection of bacteria enhancing the growth rate of axenically hatched rotifers (Brachionus plicatilis). Aquaculture 176:195-207.
Sawabe, T., I. Sugimura, M. Ohtsuka, K. Nakano, K. Tajima, Y. Ezura & R. Christen. 1998. Vibrio halioticoli sp. nov., a non-motile alginolytic marine bacterium isolated from the gut of the abalone Haliotis discus hannai. Int. J. Syst. Bacteriol. 48:573-580.
Searcy-Bernal, R. & C. Anguiano-Beltran. 1998. Optimizing the concentration of Gamma-Aminobutiric Acid (GABA) for inducing larval metamorphosis in red abalone Haliotis rufescens (Mollusca:Gastropoda). J. World Aquacult. Soc. 29:463-470.
Searcy-Bernal, R., A. E. Salas-Garza & R. A. Flores-Aguilar. 1992. Research in Mexico on the critical stage of abalone (Haliotis spp.) seed production. In: S. A. Shepherd, Tegner M. J & S. A. Guzman del Proo, (editors.). Abalone of the world: biology, fisheries and culture. Oxford: Fishing News Books. pp. 547-560.
Searcy-Bernal, R., L. A. Velez-Espino & C. Anguiano-Beltran. 2001. Effect of biofilm density on grazing and growth rates of Haliotis fulgens postlarvae. J. Shellfish Res. 20:587-591.
Son, R., G. Rusul, A. M. Sahilah, A. Zainuri, A. R. Raha & I. Salmah. 1997. Antibiotic resistance and plasmid profile of Aeromonas hydrophila isolates from cultured fish telapia (Telapia mossambica). Let. Appl. Microbiol. 24:479-482.
Takami, H., T. Kawamura & Y. Yamashita. 2002. Effects of delayed metamorphosis on larval competence, and postlarval survival and growth of abalone Haliotis discus hannai. Aquaculture 213:311-322.
Tanaka, R., I. Sugimura, T. Sawabe, M. Yoshimizu & Y. Ezura. 2003. Gut microflora of abalone Haliotis discus hannai in culture changes coincident with a change in diet. Fish. Sci. 69: 951-958.
Torkildsen, L., O. B. Samuelsen, B. T. Lunestad & O. Bergeh. 2000. Minimum inhibitory concentrations of chloramphenicol, florfenicol, trimethoprim/sufadiazine and flumequine in seawater of bacteria associated with scallops (Pecten maximus) larvae. Aquaculture 185:1-12.
Zar, J. H. 1999. Bioestatistical analysis. New Jersey: Prentice Hall. 663pp.
CASANDRA ANGUIANO-BELTRAN * AND RICARDO SEARCY-BERNAL (1)
Instituto de Investigaciones Oceanologicas, Universidad Autonoma de Baja California, Apartado Postal 453, Ensenada, B.C. 22860, Mexico
* Corresponding author. E-mail: email@example.com