Ariana Casey

BIOL 342 — Microbiology

Introduction

The human race predominantly lives on land and breathes in the air surrounding them, not giving much thought to the bodies of water that cover the majority of the earth’s surface.   In all these environments exists a common organism — bacteria.   One must look past the human —past what the naked eye can see- to understand the bacteria.   Many species exist, inhabiting a multitude of niches in the world, from the inside of soil clumps and the bottom of the ocean —devoid of oxygen- to the skin on our hands and the surfaces of plants —with aerobic respiration to sustain them.

With so many different species of microorganisms, it would be a shame to disregard the ones that live in water simply because it’s not the environment that people exist in.   With the human body and the surrounding environment being so dependent on water, is would make sense to examine this substance to find what microbes exist there.   Among one of the many uses for water that people have, the one that seemed it would have a good measure of microbiota was a fish tank filter.

The objective of the following research was to isolate a bacteria from a sampled location and to determine through physiological and genetic testing what bacteria was isolated from the sampling location.  Being that a fish tank seems a rich environment for microbiota, it is hypothesized that the bacteria will be unique, with the ability to live aerobically or anaerobically.

Methods

Sampling

The chosen location for sampling was the filter of a fish tank.   To properly sample, the filter was first removed from the water.   After removal from the fish tank, the filter was swabbed with a cotton swab, which was then streaked over the surface of the TSA plate (Lab Handout 1).   The plate was kept at room temperature for six days before being brought into lab.   At this point, a colony was pulled with a sterilized metal loop and put onto another TSA plate using a quad streak (Lab Handout 2).   As the bacteria grew, individual colonies were used to further isolate the bacteria through additional quad streaks.

At the fourth quad streak, the culture was determined to be fairly pure through microscopic observation (Lab Handout 4).  At this time, the culture was deemed pure enough to be physiologically and genetically tested to determine bacterial strain.

Physiological

One objective of obtaining a pure culture through quad streaking was to observe the colony morphology of the cells in isolate (Lab Handout 2).  To determine cell size, shape, arrangement, and wall-type, gram-staining was performed (Lab Handout 4).     Fluid thioglycollate test was performed and incubated at room temperature to determine oxygen class (Lab Handout 6).   Presence of cytochrome C oxidase was tested using an oxidase test strip with an indicator dye (Lab Handout 6).   A catalase test was used to determine if the strain of bacteria has the enzyme catalase (Lab Handout 6).   To determine which sugars the bacteria could ferment, as well as to determine other physiological characteristics which could help to characterize the isolated bacteria, an API 20E test strip was used (Lab Handout 6).   Later, due to the results with API 20E being semi-inconclusive, and with more information on the isolated bacteria, the API staph strip was used to test the bacteria.

To determine if the isolated bacteria had the ability to grow in the presence of methylene blue, and further to see if it could ferment lactose and/or sucrose, the isolate was plated onto Eosin Methylene Blue (EMB), following lab 6 protocol.   For determining if the isolate had the ability to grow in the presence of crystal violet and bile salts, and to further determine capability to ferment lactose, the isolate was plated on MacConkey (MAC) Agar (Lab Handout 6).

Additionally, to further determine the identity of the isolated bacteria, its susceptibility or resistance to various antibiotics was tested via the protocol of Lab Handout 9.  The isolated bacteria was suspended in TSB and then plated with a cotton swab onto two TSA plates, onto which antibiotic discs were placed evenly apart.  To determine susceptibility to antibiotics, zone diameter was measured once a bacterial lawn had time to form (approximately 48 hours past inoculation).  The tested antibiotics were Amakicin, Cefaperazone, Vancomycin, Trimethoprim, Oxacillin, Clindamycine, Piperacillin, and Gentamicin.

Genetic

An isolated pure culture bacteria colony was suspended into tryptic soy broth (TSB).   After a week, this TSB suspension of bacteria was used for genomic DNA extraction (Lab Handout 5).     DNA extraction was done using the PowerSoil DNA kit (MoBio), which was sequenced using Illumina MiSeq technology at the University of Alaska Fairbanks’s DNA Core Lab (Lab Handout 5).   The data procured from the Illumina Miseq sequence analysis will then be computed using BaseSpace (Lab Handout 7).  Assembly of the genome was done using the SPAdes Genome Assembler application (Lab Handout 7).  Using the Kraken Metagenomics application and the assembled genome, a taxonomic classification of the bacterial isolate was determined (Lab Handout 7).  The final application used on the data procured from the DNA Core Lab sequencing was the Prokka Genome Annotation to determine potential functional genes of the genes (Lab Handout 7).

To further analyze the results, the first node of the contigs.fasta file was pulled from the SPAdes Genome Assembly application and ran through the Nucleotide BLAST (Basic Local Alignment Search Tool) application provided by the U.S. National Library of Medicine.

Results

Physiological

The pure, isolated bacteria grew in yellow-pigmented, shiny, smooth, small, and roung colonies.  Gram-staining revealed the bacteria to Gram-positive in nature, with cocci approximately 1.5 um in diameter.  The cells were usually found in tetrads with the smallest aggregations being diplococci, though often the cells would clump further.   The fluid thioglycollate test determined the bacteria to be an obligate aerobe.   Hydrogen peroxide resulted in formation of bubbles, testing positive on the catalase test.   The oxidase test strip turned a light grey-purple in the presence of the bacterial suspension, testing positive on the oxidase test.   The API 20E tests came back predominantly negative, with the only reliable positive being the test for presence of gelatinase.   Additionally, the VP turned a slight pink within the 10 minutes after VP1 and VP2 reagents were both added.  This was inconclusive as to whether the test was positive or negative.  With the API Staph strip, there were also all negatives within 48 hours of starting the strip.   After addition of the VP and NIT reagents (there was no access to PAL reagents, so that test is inconclusive), the VP turned slight pink after the 10 minutes.  Using apiweb as a resource, I determined this to be a positive reaction.

Antibiotic testing results (Table 1) reveal this isolate to be fairly susceptible to antibiotics.

Table 1: Antibiotic Zone Diameters and Susceptibility
Antibiotic	Zone Diameter (mm)	Susceptible, Intermediate, or Resistant
Amakicin	*radius: 22	Susceptible
Cefaperazone	*radius: 18	Susceptible
Vancomycin	29	Susceptible
Trimethoprim	15	Intermediate
Oxacillin	15**	Resistant**
Clindamycine	36	Susceptible
Piperacillin	50	Susceptible
Gentamicin	36	Susceptible
*Radius was used where the zones overlapped and the diameter was impossible to measure due to overlap.  The radius of each zone was greater than the determined diameter susceptibility range. **Plated oxacillin disc was contaminated with a piece of agar from another lab mate’s bacterial isolate, and so would be inconclusive, but my bacterial isolate was used by another student that lost their isolate in a lab clean-up.  This student also used the Oxacillin disc and had a zone diameter in the resistance range.

 

Genetic

Basespace results indicate that the bacteria is likely Micrococcus luteus NCTC 2655 with 75.83% of the reads classified and 98.59% of the 73.91% analyzed reads were classified to the species level.   This is further supported by the BLAST results, which indicated a 97% identity match to Micrococcus luteus NCTC 2665.   The SPAdes Genome Assembler data revealed a genomic length of 2,615,526 base pairs with a GC content of 72.41%.   The bacteria also had 53 tRNAs and 2365 coding regions.   Probable functional genes were discovered through use of the Prokka Genome Annotation application.  Putative acetyl Co-A acetyltransferase, putative oxidoreductase, and NADH dehydrogenase-like protein SA0941 were all probable functional genes found in the genome of the isolate.

Discussion

The bacteria isolated from the fish tank filter was not a unique microbe, as was expected, but a ubiquitous microbe found in many niches.  Additionally, the bacteria was surprisingly an obligate aerobe, which was unexpected as with an environment comprised mainly of water.  The ability of the bacteria to survive in such an environment may be due to the ability of M. luteus to become quite dormant and awaken with the use of the Rpf gene (Mukamolova et al 2002; Young et al 2010).

Identification of the bacterial isolate through the genetic data alone is alright in cases where the said bacteria cannot be plated and isolated.  In the case of this study, looking at both the physiological and genetic aspects of the bacteria for being able to identify the bacteria as Micrococcus luteus NCTC 2665 strain is important.

Physiological

Micrococcus luteus NCTC 2665 “Fleming Strain’ have a distinct yellow-pigmentation to them, likely an adaptation to protect against radiation (Young et al 2010).  The bacteria is quite ubiquitous, in that it is usually found on mammalian skin, but can also be found in soil as well as fresh and marine water (Akayli et al 2016; Rickard et al 2002; Young et al 2010).  This aligns well with the sampling location being a fish tank filter, as the bacteria are found in water, plus the filter is something often handled by people, thus either could have been the source of the M. luteus at this particular location.  The bacteria is known to be coccoid, gram-positive, and form mainly tetrads (Akayli et al 2016).  With the tetrads, clumping happens, which makes sense as M. luteus are observed to create coaggregates in fresh water with some other bacteria to form a biofilm (Rickard et al 2002).  Additionally, M. luteus tests positive for both catalase and oxidase, all of which agrees with the testing done for the isolated bacteria.

The result of being an obligate aerobe from the fluid thioglycollate test also falls in line with the literature on M. luteus (Young et al 2010).  The physical test results of the API Staph test, when placed into the apiweb resource, come back with a 99.1% identity as a Micrococcus species, with an oxidase positive and yellow colony morphology leaning the test results towards Micrococcus luteus.

Genetic

With both the BaseSpace and BLAST results identifying the strain as M. luteus, especially with all the physiological evidence, it would be hard to refute that the isolated strain is, in fact, M. luteus NCTC 2665.  Beyond this, additional genetic evidence points to the identification of the strain obtained as M. luteus.  For one, the GC content of the isolated strain was 72.41%, while it is 73% for NCTC 2665 (Young et al 2010).  Strain NCTC 2665 has 2,501,097 bp, encodes 2,403 (of 2,458 total) genes to proteins, and has 48 tRNAs, which is all comparable to the isolated bacteria’s 2,615,526 bp, 2,365 coding genes, and 53tRNAs (Young et al 2010).  Of the genes, the oxidoreductase, CoA, and NADH-like protein mentioned beforehand are all important to M. luteus.  Cytochrome c oxidoreductase is a component of the respiratory chain and CoA is part of the citric acid cycle (Young et al 2010).

In discussing genes, it is important to relate some of them to physiological aspects that are seen.  A lack of glucokinase, for instance, results in M. luteus being unable to grow with glucose alone as a carbon source, which is what we witnessed through the API test strips having negative results for fermentation of glucose (Young et al 2010).  Additionally, the high susceptibility of M. luteus to antibiotics is possibly due to lack of the gene wblC, as it is a pleiotropic regulator in other bacteria that helps with resistance towards antibiotics (Young et al 2010).

Additional Discussion

The strain represented in this research is the “Fleming strain’ NCTC 2665, which is still known as Micrococcus luteus, but another strain of M. luteus —primarily ATCC 9341- has been reclassified as Kocuria rhizophila (Tang and Gillevet 2003).  In fact, the NCTC 2665 strain had, “recently’ (in accordance to a 2010 paper) been declared taxonomically separate from K. rhizophila (Young et al 2010).

Strain ATCC 9341 is different in a few ways, some being that it is oxidase negative compared to the oxidase positive nature of NCTC 2665, as well as that the strain ATCC 9431 forms acids from glucose and fructose, neither of which were positive tests on the API test strips performed, meaning that my bacteria isolate does not confer with the traits presented by the Micrococcus luteus ATCC 9431 strain that was reclassified as Kocuria rhizophila (Tang and Gillevet 2003).  It, indeed, appears that the isolated bacteria follows more closely with the “Fleming strain’ NCTC 2655 of M. luteus which has not been reclassified to Kocuria rhizophila.

 

References

Akayli, T., Albayrak, G., Ürkü, Ç, Çanak, Ö, & Yörük, E. (2015). Characterization of Micrococcus luteus and Bacillus marisflavi Recovered from Common Dentex (Dentex dentex) Larviculture System. Mediterranean Marine Science, 17(1). doi:10.12681/mms.1322

Mukamolova, G. V., Turapov, O. A., Young, D. I., Kaprelyants, A. S., Kell, D. B., & Young, M. (2002). A family of autocrine growth factors in Mycobacterium tuberculosis. Molecular Microbiology, 46(3), 623-635. doi:10.1046/j.1365-2958.2002.03184.x

Rickard, A., McBain, A., Ledder, R., Handley, P., & Gilbert, P. (2003). Coaggregation between freshwater bacteria within biofilm and planktonic communities. FEMS Microbiology Letters, 220(1), 133. doi:10.1016/S0378-1097(03)00094-6

Tang, J. S. and Gillevet, P. M. (2003). Reclassification of ATCC 9341 from Micrococcus luteus to Kocuria rhizophila. International Journal Of Systematic And Evolutionary Microbiology, 53(4), 995-997. doi:10.1099/ijs.0.02372-0

Young, M., Artsatbanov, V., Beller, H. R., Chandra, G., Chater, K. F., Dover, L. G., . . . Greenblatt, C. L. (2009). Genome Sequence of the Fleming Strain of Micrococcus luteus, a Simple Free-Living Actinobacterium. Journal of Bacteriology, 192(3), 841-860. doi:10.1128/jb.01254-09

Micrococcus luteus in a Veterinary Clinical Setting

Erin Murray

Biol 342, Section F02

 

Introduction

In veterinary clinics or hospitals, it is highly important to maintain as clean an environment as possible for the sake of their patients’ health. Failure to do so carries the possibility of inadvertently transferring bacterial or viral pathogens from one patient to the next, which could result in a healthy patient becoming sick simply from having visited the clinic. This is known as a nosocomial infection (Stull and Weese). If a patient is hospitalized and acquires a nosocomial infection, this could lead to its hospital stay being prolonged (Harper et al.). Microbes known to cause nosocomial infections have been isolated from samples taken from various surfaces around veterinary clinics, including medical equipment used on patients, as well as from air samples from the various rooms of an animal hospital (Harper et al.). This means that airborne bacteria could potentially land on and colonize surfaces of medical equipment even after they have been thoroughly cleaned.

In the study of airborne sampling, the most commonly isolated bacteria belonged to the genus Micrococcus (Harper et al.). Micrococcus species commonly grow on Mammalian skin as a commensal bacteria, and are rarely ever pathogenic. There have been rare cases where patients developed infections from Micrococcus bacteria becoming an opportunistic pathogen, but this has only occurred in patients that were immunocompromised (Hanafy et al., Public Health England).

The purpose of this study was to attempt to determine if any pathogenic bacteria were taking up residence on the surfaces of the medical equipment at a local emergency veterinary clinic. Speaking from personal experience, one potential source of contact transfer of microbes is a veterinarian’s or technician’s stethoscope. Though it is used on every patient walking through the doors of the clinic, the cleaning of this piece of medical equipment can be easily overlooked. Many veterinary staff will wear their stethoscope around their neck in order to keep it close at hand and may not get a chance to, or may even forget to, disinfect it in between patients.

I hypothesized that I would find species of bacteria that typically lived either on Mammalian skin and hair, or in canine or feline mouths. I suspected that I might find mouth bacteria because dogs and cats lick their fur to groom themselves. From this hypothesis I predicted that I would isolate a bacterium that could at least tolerate oxygen and would therefore be an aerobe or an aerotolerant anaerobe.

 

Methods

The diaphragm of a frequently used stethoscope at a local veterinary clinic was swabbed with a sterile cotton swab that had been wetted with sterile water. The swab was then used to inoculate a Tryptic Soy Agar (TSA) plate. The plate was sealed with parafilm and was stored at room temperature for one week to allow for bacterial colonies to grow. At the end of one week, several colonies of different colors and textures had grown, and one colony was selected to be isolated. The colony was goldenrod yellow in color, circular and raised, had a smooth and shiny finish, and was about the size of a pencil eraser. A sample of the colony was transferred onto a new TSA plate using the quadrant streak technique as outlined in the handout for Lab 2: Aseptic Technique. The plate was incubated at 37 °C to accelerate colony growth. A new quadrant streak was performed every 3-4 days for a total of 4 times to ensure that purity of the culture had been achieved.

A series of physiological tests were performed on the isolate to determine its physiological characteristics. A Gram stain was performed on a microscope slide of the culture, following the protocol as outlined in Lab 4: Staining Techniques, to determine the type of cell wall the bacteria possessed. Following the protocols as outlined in Lab 6: Physiological Testing of Your Isolate, a fluid thioglycollate test was performed to determine the culture’s oxygen class, an oxidase test was performed to determine whether the isolate produced the cytochrome c oxidase enzyme, and a catalase test was performed to determine whether the isolate produced the catalase enzyme. An API 20E test strip was performed, following the protocol as outlined in Lab 6, to determine which materials the bacteria could metabolize. The test strip was rechecked after 24-48 hours of incubation at 37 °C. Later on, an API Staph test strip was also performed, following the protocol on the API Staph handout. It, too, was incubated at 37 °C and rechecked 24-48 hours later.

The culture’s susceptibility and/or resistance to various antibiotics was tested, following the protocol as outlined in Lab 9. The antibiotics used for the test were amikacin, cefazolin, clindamycin, erythromycin, gentamicin, oxacillin, tetracycline, and tobramycin.

DNA was extracted from the isolate for genomic analysis, using the PowerSoil DNA isolation kit and following the protocol as outlined in the handout for Lab 5. The DNA sample was then submitted to the UAF DNA Core Lab to be sequenced. When the raw genomic data had been obtained, it was then analyzed on BaseSpace, using the SPAdes Genome Assembler, Kraken Metagenomics, and Prokka Genome Annotation programs. The contig data was then analyzed using the NCBI Nucleotide BLAST program.

 

Results

Gram staining of the bacterium revealed that the isolate was a Gram-positive cocci that formed tetrads and irregular clusters. In fluid thioglycollate the bacteria grew directly on and just below the surface of the fluid, and no growth was observed in the anoxic layer of the fluid, therefore the bacterium is likely a strict anaerobe. The isolate tested positive for the presence of the enzymes catalase and cytochrome c oxidase. The API 20E and API Staph assays had no positive results. Antibiotic resistance testing revealed that the isolate was susceptible to amikacin, cefazolin, clindamycin, gentamicin, tetracycline, and tobramycin, was intermediately susceptible to oxacillin, and was resistant to erythromycin.

SPAdes Genome Assembler discovered 300 contigs with the largest contig being 61,411 base pairs (bp). The total length of the genome was determined to be 2.51 Mbp, and the GC content of the genome was determined to be 72.72%. Prokka Genome Annotation discovered 53 tRNAs, 0 rRNAs, 1 CRISPR gene, and 2270 CDs. The Kraken Metagenomics program took 173,940 reads. Of those reads, 55,285 were classified, resulting in 31.78% of total reads being classified. 25.85% of the total reads were classified to the species level, and the bacterium was identified as Micrococcus luteus. Analyzing contigs via BLAST revealed that the sequence from node 21 was a 97% match to the genomes of the NCTC 2665 and trpE16 strains of M. luteus.

 

Discussion

After extensive testing of the isolate, it has been concluded that the identity of the isolate is Micrococcus luteus.

M. luteus is described in the literature as an aerobic, catalase- and oxidase-positive, Gram-positive non-motile cocci which form tetrads, with colonies being yellow in pigment, convex and smooth in texture with regular edges (Public Health England, Whitman et al.). The physical appearance of colonies of the isolate grown on agar plates, and physiological tests performed on the isolate are in agreement with these descriptions. Genome length and GC content of the isolate that were reported in SPAdes also agree with values in the literature (Hanafy et al., Whitman et al.)

Neither of the API assays that were attempted had any color changes, meaning that none of the tests came back positive. For the API 20E test, the lack of positive results is likely due to the test being the wrong fit for this microbe – it was meant to be used on Gram-negative anaerobes, and the isolate was a Gram-positive strict aerobe. It is possible that the reagents were unable to penetrate the thick peptidoglycan layer of the bacteria’s cell walls in order for metabolism of the reagents to occur.

The API Staph test should have been the best fit for the isolate. However, it still came back with no positive results. Researching the literature reveals that M. luteus should be able to metabolize glucose, fructose, sucrose, and mannose (Hanafy et al.), meaning that the GLU, FRU, SAC, and MNE wells on the API Staph test should have had color changes to indicate positive results. Some strains have been found to also metabolize maltose and trehalose, contrary to prior data indicating that these molecules were not utilized by M. luteus (Wieser et al.), meaning that there possibly could have been positive results from the MAL and TRE wells if the isolate was a similar strain. The literature also indicates that M. luteus is urease positive (Whitman et al.), so the URE well should have indicated a positive result as well. The lack of results from the API Staph test was probably due to not having a fresh enough sample: the instructions for the test call for a sample that is 18-24 hours old, and the plate that the sample was taken from had been incubating for between 4 and 7 days.

The isolate was found to be susceptible to most of the antibiotics that were tested, except for erythromycin, which it was completely resistant to. Resistance to erythromycin has been found to be due to a plasmid present in a strain of M. luteus isolated from human skin (Liebl et al.). With humans and companion animals having such an intimate relationship and being in contact with each other so often, it doesn’t take a stretch of the imagination to suggest that erythromycin-resistant M. luteus could be transferred from human skin onto the skin of a companion animal, or vice versa, which would explain the presence of erythromycin-resistant M. luteus on a stethoscope in a veterinary clinic. Horizontal gene transfer of this resistance plasmid could potentially cause other bacterial species to develop resistance as well.

Further investigation into the types of bacteria growing on medical equipment in veterinary clinics should be pursued. Only one of several different colonies from the original swab of the stethoscope was chosen for further evaluation. Analysis of the other colonies that were present on the original TSA plate would be the first step in continuing the investigation, followed by taking samples from other objects and equipment, and possibly obtaining airborne samples from around the clinic. This type of investigation would be very helpful in assessing local clinics’ risk of exposing their patients to nosocomial infections.

 

Figures

Figure 1. Macroscopic and microscopic morphology of the isolate. Colonies are yellow, convex, and smooth in texture with a regular edge. Gram staining reveals that the bacteria are Gram-positive cocci that form tetrads and irregular clusters.

Figure 2. Antibiotic resistance testing of the isolate. The isolate showed resistance to erythromycin, but to no other antibiotic which was tested.

Figure 3.  Krona Classification chart from the Kraken Metagenomics program. The reads were classified and nested into successive taxonomic groups, down to the species level.

 

References

Hanafy R.A., Couger M.B., Baker K., Murphy C., O’Kane S.D., Budd C., French D.P., Hoff W.D., and Youssef N. 2016. Draft genome sequence of Micrococcus luteus strain O’Kane implicates metabolic versatility and the potential to degrade polyhydroxybutyrates. Genomics Data 9:148—153.

Harper T.A., Bridgewater S., Brown L., Pow-Brown P., Stewart-Johnson A., Adesiyun A.A. 2013. Bioaerosol sampling for airborne bacteria in a small animal veterinary teaching hospital. Infection Ecology and Epidemiology 3.1.

Liebl W., Kloos W.E., and Ludwig W. 2002. Plasmid-borne macrolide resistance in Micrococcus luteus. Microbiology 148:2479—2487.

Public Health England 2014. Identification of Staphylococcus species, Micrococcus species and Rothia species. Bacteriology — Identification 3:1-32.

Stull J.W., and Weese J.S. 2015. Hospital-Associated Infections in Small Animal Practice. Veterinary Clinics: Small Animal Practice 45:217—233.

Whitman M., Goodfellow M., Kämpfer P., Busse H.J., Trujillo M., Ludwig W., Suzuki K.I. 2012. Bergey’s Manual of Systematic Bacteriology: Volume 5: The Actinobacteria. Springer Science & Business Media, p. 574-575.

Wieser M., Denner E.B.M., Kämpfer P., Schumann P., Tindall B., Steiner U., Vybiral D., Lubitz W., Maszenan A.M., Patel B.K.C., Seviour R.J., Radax C., and Busse H.J. 2002.  Emended descriptions of the genus Micrococcus, Micrococcus luteus (Cohn 1872) and Micrococcus lylae (Kloos et al. 1974). International Journal of Systematic and Evolutionary Microbiology 52:629—637.

 

Microbes are an omnipresent living force found in every environment and habitat on the planet, from deep below the ocean in the crushing darkness at hydrothermal vents, to within human bodies and floating in the air around us. The diversity of microbes, especially bacteria, is staggering and constantly shifting through the tremendous evolutionary prowess and genetic flexibility of these organisms.

For the purposes of this research, I chose to investigate the microfauna of my aquarium biome. I have worked to culture a realistic and comfortable habitat for the organisms I keep, which has involved obtaining live plants, cultivating algae, using a heater to maintain constant temperature, and using a LED lamp to mimic sunlight. Nitrogen fixing and ammonia-oxidizing bacteria are necessary for a healthy aquarium environment (Urakawa et al., 2008)(Zehr et al., 2000), and bacteria considered pathogenic to fish and aquatic plants are perpetually present in the water (Baran et al., 2008)(Smith et al., 2012).

My goal was to isolate and identify a bacterial strain from my aquatic microbiome, then determine the likely role it plays within the system I have worked to cultivate. To do so, I sampled the aquarium water, allowed the bacteria to grow colonies from which to extract pure culture, and isolate a pure culture from which to run both genomic and physiological tests upon. I theorized the microbe I isolated is a form of nitrogen-fixing bacteria, as my tank is a healthy system with no diseased fish or plants, and has been thriving with minimal water changes, a sign of nitrogen cycling within the ecosystem.

However, the microbe I identified was Rhodococcus erythropolis, a versatile and occasionally pathogenic bacteria associated with aquatic and soil environments. This was somewhat surprising due to the predicted relative rarity of R. erythropolis in comparison to other microbial populations, such as nitrogen fixers in aquatic environments, but considering the methods I used, the bacteria I chose had already been through a selective process once I decided to isolate.

 

Methods

Sampling and Growth

The aquarium biome was sampled by submerging a sterile swab of cotton into the water and gently swiping it across a Betta splendens (Betta fish). The swab was then wiped across a plate of sterilized SA agar and left to incubate at room temperature for 6 days. Bacterial growth appeared after 72 hours in three small colonies. The SA plate had initially been intended to select for fungal growth, but I chose to isolate and study the acidophilic bacteria growing on the plate instead.

These bacterial colonies were isolated into pure culture by the quadrant streak method (Lab 2 Handout). Four successive streak plates were made over the course of 3 weeks, and were incubated at 37 °C during the growth phase of the bacterium.

 

Identification

When the isolate was sufficiently purified, a Gram stain was conducted to determine if the strain is Gram-positive or Gram-negative. This test was conducted according to the methods of Lab Handout 4.

The isolate was also tested for a variety of physiological abilities. A fluid thioglycollate test was used to determine the strain’s oxygen class, a catalase test establishes whether a bacterium contains the catalase enzyme, and a oxidase test to see if the isolate contained cytochrome c oxidase. An API 20E test strip was additionally used to test the isolate’s capacity for fermentation of various sugars, the presence of specific enzymes, and decarboxylations of amino acids, with 20 tests in total. All physiological tests were completed according to the methods of Lab Handout 6.

To obtain a more specific idea of the bacterium’s identification, DNA extraction and whole genome sequencing was conducted on the organism. Extraction was completed with a freshly streaked isolate and the PowerSoil Isolation kit according to the methods of Lab Handout 5. Genomic sequencing was completed in the Illumina MiSeq DNA sequencer at the University of Alaska-Fairbank’s DNA Core Lab, and analyzed in BaseSpace with SPAdes Genome Assembler and Kraken metagenomics according to the instructions of Lab Handout 6. These tests would determine the closest genetic match to the extracted DNA, and how many contigs, functional genes, and tRNA regions are identified, as well as what each section coded for, when relevant.

Unfortunately, after genomic extraction, the original isolate was lost. I attempted to reisolate the same bacteria again through similar sampling methods and selection of a colony of visually identical microbes, but was unsuccessful. Therefore, any antibiotic resistance by my original strain remains unknown, as well as its ability to ferment lactose or sugars, as would have been tested in Labs 8 and 9.

 

Results

This isolate was observed to grow most quickly at 25 °C. Colonies were matte white and circular with defined edges and a convex shape. This bacteria was able to grow under both refrigeration and at 37 °C, though slowly.

Gram staining of the isolate showed the microbe as a gram positive bacillus. The rods most commonly appeared clustered in pairs (diplobacilli), bent at an angle so as from a distance they appear to be one spiral bacteria. The individual bacilli are approximately 0.5μm long.

 

Figure 1. Isolate gram-stained and magnified at 1000X. This organism is clearly gram-positive and is clustered in pairs with few exceptions.

 

In physiological tests, the isolate was positive in the catalase and oxidate tests. (Acharya, 2015). Complete results for the API 20E test strip are listed in Table 1. The ONPG test (tests for β-galactosidase enzyme) was inconclusive due to an unfixable air bubble that hindered results, and is not included.

 

Test	Result
ADH- determines decarboxylation of the amino acid arginine by arginine dihydrolase	Negative
LDC- determines decarboxylations of the amino acid lysine by lysine decarboxylase	Negative
ODC- determines decarboxylations of the amino acid ornithine by ornithine decarboxylase	Negative
CIT- detects utilization of citrate as only carbon source	Negative
H2S- detects production of hydrogen sulfide	Negative
URE- detection of enzyme urease	Positive
TDA- detection of the enzyme tryptophan deaminase	Positive
IND- detection of production of indole by the enzyme tryptophanase	Positive
VP- determines if fermentation of glucose by bacteria utilizing the butylene glycol pathway is being utilized	Positive
GEL- tests for the production of the enzyme gelatinase	Negative
GLU- detects fermentation of glucose (a hexose sugar)	Positive
MAN- detects fermentation of mannose (a hexose sugar)	Positive
INO- detects fermentation of inositol (a cyclic polyalcohol)	Positive
SOR- detects  fermentation of sorbitol (a alcohol sugar)	Positive
RHA- detects fermentation of rhamnose (a methyl pentose sugar)	Positive
SAC- detects fermentation of sucrose (a disaccharide)	Negative
MEL- detects fermentation of melibiose (a disaccharide)	Negative
AMY- detects fermentation of amygdalin (a glycoside)	Negative
ARA- detects fermentation of arabinose (a pentose sugar)	Negative

Table 1. A complete listing of the results from my API 20E test strip for my isolate (Acharya, 2015).

 

In the fluid thioglycollate test, the isolate proved able to survive in all areas of oxygenation, but preferred to grow at the surface, making it a facultative aerobe. Growth was thickly concentrated at the surface and directly below the surface of the agarose, but a thin line of microbial growth was visible extending from the surface to the base of the tube.

Genomic tests indicated only approximately 70% of genetic data was usable, as the other data was not even classified as bacteria in the analysis. However, within the usable information, the confidence level of the identified sequences were 95% or greater. Complete percentages of taxonomic rank are illustrated in Table 2, indicating a high level of precision in the identification of the isolate as Rhodococcus erythropolis (BaseSpace, 2017).

 

Kingdom	Phylum	Class	Order	Family	Genus	Species
Bacteria	Actinobacteria	Actinobacteria	Actinomycetales	Nocardiceae	Rhodococcus	erythropolis
99.11%	98.41%	98.41%	98.34%	95.99%	96.05%	95.91%

Table 2. The confidence levels of identified sequences within the bacterial genome and the identified classifications. Note all of these values are derived from ∽70% of total data, but the remaining 30%  of data was completely unidentifiable, even to the kingdom level, and as such has been disregarded in this interpretation.

 

Only one contig of 1420bp was identified. Additionally, only one tRNA was observed, and 169 coding regions.

Lastly, BLAST analysis was used as a secondary classification system to check the accuracy of results from Kraken metagenomics. In BLAST, the result for nucleotide analysis on contigs came back as 97-94% match for R. erythropolis, confirming the Kraken results (National Center for Biotechnology Information, 2017).

 

Discussion

The results of the genomic tests led me to believe the strain I isolated was a variety of R. erythropolis, despite having relatively small amounts of genetic data to work with. The lack of contigs, functional genes, and tRNA leaves room for there to be a considerable amount of error in the identification, but the certainty of identification for what is present from the sample is considerably confident.

Additionally, if we are to analyze the characteristics of R. erythropolis, it is a Gram-positive, mesophilic, aerobic rod bacteria. Interestingly, it was recently observed to be a human pathogen under specific circumstances, causing septicemia in immunocompromised individuals (Park et al., 2011). Ironically, a fish in the tank I sampled died of septicemia earlier this year, though there is no means to determine if R. erythropolis was responsible for the death.

These organisms are commonly found in both soil and aquatic environments, as well as eukaryotic cells. In wild environments, they are known for utilizing a wide variety of organic compounds. Different strains have demonstrated the ability to undergo oxidations, dehydrogenations, epoxidations, hydrolysis, hydroxylations, dehalogenations and desulfurization in order to process the resources of their habitat (de Carvalho, 2005). Due to this diversity, Rhodococcus sp. are considered important industrial organisms, utilized in production of bioactive steroids and fossil fuel biodesulfurization, and is also regarded as the most commercially successful microbial biocatalyst (McLeod, 2006).

Comparatively, my isolate demonstrated several traits associated with Rhodococcus species. My isolate was gram-positive and grew in rods. The diplobacilli arrangement noted in my isolate was never mentioned as a distinguishing factor of R. erythropolis or other Rhodococcus species. I collected my sample from an aquatic environment, and though the environment I collected it from was warm, I found the bacterium grew much more successfully at room temperature. Additionally, my bacteria was established as tolerant of acidic environments from the initiation of the experiment. R. erythropolis is not characterized as an acidophile, so I believe it is an ability it possesses, but as demonstrated by growth on TSA plates and TSB, not the preferred environment of the organism.

Physiological tests revealed my bacteria sample can process urea, produces the enzymes tryptophan deaminase and tryptophanase, and can ferment a variety of simple sugars, but does not appear to have the dramatic abilities some R. erythropolis strains, though some were never tested. Although my isolate thrived in an aerobic environment, a characteristic of Rhodococcus sp., it also showed the capacity to persist in all oxygen zones, something not all Rhodococcus sp. can do.

Due to the genomic and physical evidence presented, I believe my isolate is a variety of Rhodococcus, though it may be debatable whether it is R. erythropolis, considering the lack of additional genomic data, and the diversity of the Rhodococcus genus. It is commonly acknowledged that Rhodococcus is environmentally widespread, and many more species may be present than currently documented in journals or databases (Park, 2011).

Several tests were conducted in lab research on other isolates which were not possible to do on my bacterium, due to its unfortunate and premature loss from the lab samples. As such, there can be no further research on this specific organism, though if there were, would like to investigate its abilities in several other ways.

I especially would have liked to analyze antibiotic resistance of my isolate. As it is an environmental microbe, especially one which can compete in the same pH zone as fungi, I would imagine it would have resistance to fungal antibacterials so as to better contend with fungi living in the same niche. Additionally, since it has potential to be pathogenic, it would have been beneficial to see what resistance or susceptibility it may have.

Though it would have been interesting and insightful to test my isolate’s ability to process petroleum, considering the use of R. erythropolis as a biodegradation, this was an opportunity not presented to me in this study. Another experiment I would have liked to complete in future research is analysis of the pathogenic potential of my specific strain. As there are multiple accounts of R. erythropolis causing septicemia in patients, perhaps analyzing growth on blood agar would show how quickly an infection of my isolate could become threatening.

In conclusion, my bacterium sampled was not a nitrifying organism, as I had predicted. Rather, it is a decomposer with the capacity to degrade a variety of environmental compounds, likely taking full advantage of the waste products and other organic matter present in the aquarium biome I took it from. There is a wide range of further experiments I would have liked to have conducted on this organism, but due to the follies of lab, I have, for the moment, lost the ability to further study this opportunistic organism. However, this entire study was a reinforcement of the staggering amount of diversity within the microbial world and the startlingly prolific world just out of our sight at all times.

 

References

Archarya, T. (2015). API 20E Test System: Introduction, Procedure Results and Interpretations. Retrieved from https://microbeonline.com/api-20e-test-system-introduction-procedure-results-interpretations/ on April 2, 2017.

 

Baran V., Dvorska L., Matlova L., Pavlik I., Svastova P. (2008). Distribution of mycobacteria in clinically healthy ornamental fish and their aquarium environment. Journal of Fish Diseases 29: 383—393.

 

de Carvalho, C.C.C.R., da Fonseca, M.M.R. (2005). The remarkable Rhodococcus erythropolis. Applied Microbiology and Biotechnology 67: 715—726.

Illumina BaseSpace. (2017). SequenceHub. Retrieved from https://basespace.illumina.com/dashboard on April 2, 2017.

 

McLeod, M.P., et al. (2006) The complete genome of Rhodococcus sp. RHA1 provides insights into a catabolic powerhouse. Proceedings of the National Academy of Sciences of the United States of America 103: 15582—15587.

 

National Center for Biotechnology Information. (2017). BLAST Nucleotide Sequence. Retrieved from  https://blast.ncbi.nlm.nih.gov/Blast.cgi?PROGRAM=blastn&PAGE_TYPE=BlastSearch&LINK_LOC=blasthome on April 2, 2017.

 

Park, S.D., Uh, Y., Jang, I.H., Yoon, K.J., Kim, H.H., Bae, Y.J. (2011). Rhodococcus erythropolis septicaemia in a patient with acute lymphocytic leukaemia. Journal of Medical Microbiology 60: 252—255.

 

Smith K., Schmidt V., Rosen G.E., Amaral-Zettler L. (2012). Microbial Diversity and Potential Pathogens in Ornamental Fish Aquarium Water. PLOS One: https://dx.doi.org/10.1371/journal.pone.0039971.

Urakawa H., Tajima Y., Numata Y., Tsuneda S. (2008). Low Temperature Decreases the Phylogenetic Diversity of Ammonia-Oxidizing Archaea and Bacteria in Aquarium Biofiltration Systems. Applied and Environmental Microbiology 74: 894-900.

 

Zehr J.P., Carpenter E.J., Villareal T.A. (2000). New perspectives on nitrogen-fixing microorganisms in tropical and subtropical oceans. Trends in Microbiology 8: 68-73.