Isolation and Purification of Planctomycetes associated with Harbor and Lagoon Seagrasses of the Red Sea

Thesis by

Holly Bream

In Partial Fulfillment of the Requirements

For the Degree of

Master of Science

King Abdullah University of Science and Technology

Thuwal, Kingdom of Saudi Arabia

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EXAMINATION COMMITTEE APPROVALS FORM

The thesis of Holly Bream is approved by the examination committee.

Committee Chairperson Dr. Vladimir Bajic

Committee Member Dr. Uli Stingl

Committee Member Dr. Pascal Saikaly

Committee Member Dr. Feras Lafi

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ABSTRACT

Isolation and Purification of Planctomycetes associated with Harbor and Lagoon Seagrasses of the Red Sea

Holly Bream

Planctomycetes are members of a unique superphylum along with Verrucomicrobia and

Chlamydia, situated in the domain Bacteria. They have distinct structural and morphological features, and discoveries made through phylogenetic analysis indicate their important role in nutrient cycling. Their known relationships with marine photosynthetic organisms led to the formation of this study's hypothesis, namely, that

Planctomycetes can be isolated from the biofilm of seagrass species of the Red Sea using cultivation techniques adapted for these organisms. Preparation of solid and liquid media using M13 with both agar and gellan, and 2216 Difco Marine Broth full- strength, 1/10-strength, and with antibiotics, resulted in the successful isolation of

Planctomycetes as confirmed by morphological examination and transmission electron microscopy. The work performed in this study provides a solid foundation for further studies to elucidate the metabolic pathways and ecological significance of

Planctomycetes.

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ACKNOWLEDGEMENTS

Many thanks to Dr. Vladimir Bajic, my committee chair, for his support of the microbiological work we were able to perform in the CBRC lab, and for patiently awaiting the arrival of data that the CBRC team can play with. Thank you to Dr. Feras

Lafi, without whom this work would not have been possible, from the conception of the project through the outline of the thesis. I would also like to thank the invaluable members of my thesis committee, Dr. Uli Stingl and Dr. Pascal Saikaly, for their insight and constructive criticism of the draft stages of this work, and for being willing to help me improve it even over the course of a very hot Saudi summer.

I much appreciate the time and effort put in by Dr. Abdulaziz Al-Suwailem and Dr. Zenon

Batang of the CMOR Core Lab. Their professionalism and knowledge greatly enhanced the collection and analysis phases of the experiment.

A special thanks goes to Jay Larson, who reminded me when it was appropriate to eat meals, shower, and occasionally take breaks during the preparation of this thesis. I may not have survived through its completion otherwise.

To the newly married Mrs. Soha Alamoudi, I would like to extend my appreciation for keeping me company during the long hours we shared under the fume hood, and until all hours of the night after the collection trips. She inspired me to work harder every day and is a model of strength and discipline.

Last but not least, I would like to thank my family. They may be 7,500 miles away physically, but were with me in spirit throughout the writing of this thesis as I missed birthdays, graduations, and weddings. I dedicate this to you. 5

TABLE OF CONTENTS

Page

EXAMINATION COMMITTEE APPROVALS FORM ...... 2

ABSTRACT ...... 3

ACKNOWLEDGEMENTS ...... 4

TABLE OF CONTENTS ...... 5

LIST OF ABBREVIATIONS ...... 6

LIST OF FIGURES ...... 9

LIST OF TABLES ...... 11

CHAPTER 1: INTRODUCTION………………………………………………………………………...... 13

CHAPTER 2: METHODOLOGY………………………………………………………………………...... 49

CHAPTER 3: RESULTS AND DISCUSSION………………………………………………………...... 64

CHAPTER 4: CONCLUSION………………………………………………………………………...... 79

REFERENCES………………………………………………………………………………………………………….. 83

APPENDICES…………………………………………………………………………………………………………... 92

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LIST OF ABBREVIATIONS

Acetyl-CoA acetyl co-enzyme A

ATP adenosine triphosphate

β-RFAP β-ribofuranosylaminobenzene 5ʹ-phosphate

B (Appendix B) broth

C1 one-carbon molecule

CCD charge-coupled device

CMOR Coastal and Marine Resources Core Lab dd H2O double-distilled water

EDTA ethylenediaminetetraacetic acid

F (Appendix B) flask

Fae formaldehyde-activating enzyme

FISH fluorescence in situ hybridization

Fmd formyl methanofuran dehydrogenase

Ftr formylmethanofuran-tetrahydromethanopterin N- formyltransferase

G (Appendix B) glycerol

H4F tetrahydrofolate

H4MPT methylene tetrahydromethanopterin

HAO hydroxylamine:oxygen oxidoreductase

HD hydrazine dehydrogenase

HH hydrazine hydrolase 7

ICM intracytoplasmic membrane

KAUST King Abdullah University of Science and Technology

LECA last eukaryotic common ancestor

LGT lateral gene transfer

LUCA last universal common ancestor

MC membrane coat

Mch methenyl-H4MPT cyclohydrolyase

MtdA methylene-H4MPT dehydrogenase A

NIR nitrite oxidoreductase

NPC nuclear pore complex

NTA nitrilotriacetate

Omp2 outer membrane protein 2

PCR polymerase chain reaction

PMF proton motive force

PVC Planctomycetes-Verrucomicrobia-Chlamydiae

PVC polyvinyl chloride (in Methodology only) redox reduction oxidation

RO research objective rpm revolutions per minute

S (Appendix B) sludge sed (Appendix B) sediment

SRSW sterilized Red Sea water 8

TEM transmission electron microscopy

TRFLP terminal restriction fragment length polymorphism

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LIST OF FIGURES

Figure 1. Location of Al Kharrar Lagoon...... 16

Figure 2. Location of Rabigh Harbor...... 19

Figure 3. C. serrulata as depicted by Seagrass-Watch…………………………………………………..23

Figure 4. H. uninervis as documented by the Systematic Marine Biodiversity Inventory System………………………………………………………………………………………………………………….………24

Figure 5. H. stipulacea as identified by DAISIE Species Factsheet…………………………….……25

Figure 6. Diagram of the cellular organization of Pirellula and Isosphaera species, and “Candidatus” Brocadia anammoxidans and Gemmata species from Fuerst...... 32

Figure 7. Schematic of possible anammoxosome membrane reactions involved in catabolism and reverse electron transport chain...... 44

Figure 8. Map of sampling sites at Al Kharrar Lagoon and Rabigh Harbor...... 57

Figure 9. Seagrass samples collected in Rabigh Harbor and Al Kharrar Lagoon with their taxonomic diagnosis displayed...... 65

Figure 10. Photo of 2216 plate inoculated with undiluted H. uninervis biofilm sample...67

Figure 11. Bacteria present in the white rosette-like flakes accompanying the ball-like structures identified in enrichment broth...... 70

Figure 12. Specimens from a colony picked on an anti-2216 plate that was inoculated with sludge swabbed from the side of a flask enriching H. stipulacea abnormal biofilm sample…………………………………...... 71

Figure 13. Specimens processed from sample grown on an M13 plate which was streaked from broth enrichment of sediment collected near H. stipulacea and observed using TEM after negative staining with uranyl acetate...... 72

Figure 14. Comparison of suspected Pirellula species with confirmed Pirellula strains……………………………………………...... 76

Figure 15. Flagella and crateriform structures clearly visible in cell from FHS-sed2 sample...... 125 10

Figure 16. Close-up of complex sheathed flagella...... 126

Figure 17. Specimen displaying polar fibrillar appendages...... 127

Figure 18. Two cells attached to each other by some unknown mechanism, both displaying polar fibrillar appendages...... 128

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LIST OF TABLES

Table 1.Rabigh Harbor and Al Kharrar Lagoon Environmental Parameters…………………..58

Table 2. Description of colonies growing from all three concentrations of H. uninervis inoculum on all five media types as of 29/4/12, with updates on isolates that were picked previously...... 94

Table 3. Description of colonies growing from all three concentrations of C. serrulata inoculum on all five media types as of 29/4/12, with updates on isolates that were picked previously………………...... 95

Table 4. Description of colonies growing from all three concentrations of “normal” H. stipulacea inoculum on all five media types as of 29/4/12, with updates on isolates that were picked previously………...... 97

Table 5. Description of colonies growing from all three concentrations of abnormal H. stipulacea inoculum on all five media types as of 29/4/12, with updates on isolates that were picked previously………...... 99

Table 6. Description of colonies growing from all three concentrations of C. serrulata sediment inoculum on all five media types as of 1/5/12……………...... 100

Table 7. Description of colonies growing from all three concentrations of H. uninervis sediment inoculum on all five media types as of 1/5/12…………………………...... 101

Table 8. Description of colonies growing from all three concentrations of H. stipulacea sediment inoculum on all five media types as of 1/5/12…………………………………..…...... 102

Table 9. First description from 1/5/12 of plates streaked using inocula from H. uninervis biofilm samples that were first enriched in selective media...... 103

Table 10. First description from 1/5/12 of plates streaked using inocula from C. serrulata biofilm samples that were first enriched in selective media...... 103

Table 11. First description from 1/5/12 of plates streaked using inocula from H. stipulacea biofilm samples that were first enriched in selective media...... 104

Table 12. First description from 1/5/12 of plates streaked using inocula from sediment samples associated with all three seagrasses that were first enriched in selective media...... 105

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Table 13. Second look on 7/5/12 at C. serrulata plates...... 106

Table 14. Second look on 7/5/12 through 9/5/12 at H. uninervis plates...... 108

Table 15. Second look on 9/5/12 and 10/5/12 at H. stipulacea abnormal plates...... 110

Table 16. Second look on 10/5/12 at H. stipulacea “normal” plates...... 112

Table 17. Second look on 11/5/12 and 12/5/12 at all plates with inocula originating from flasks...... 114

Table 18. Second look on 12/5/12 and 13/5/12 at all plates with inocula originating from sediment...... 117

Table 19. Examination of plates originating from plates inoculated with biofilm, recorded on 13/5/12 and 14/5/12...... 119

Table 20. Examination of plates originating from plates inoculated from enrichment flasks, recorded on 14/5/12...... 123

Table 21. Examination of plates originating from plates inoculated with sediment, recorded on 15/5/12...... 124

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CHAPTER 1: INTRODUCTION

The eastern coastal area of the Red Sea is dotted with hypersaline lagoons, known in

Arabic as sharms. Their biodiversity is vastly understudied, with only a small number of papers published pertaining to the sedimentological composition of the sharms (El-

Sayed 1987; Abd and Awad 1991; Ouf 1994; Washmi 1999; Basaham 2008), a handful on benthic foraminifera distribution (Hariri 2008), and, more recently, one publication on bioactive compounds isolated from bacteria living in association with eukaryotes located in Red Sea sharms (Xu, Kersten et al. 2012). The refuge these sharms offer to their inhabitants results in unique ecosystems set apart from the greater Red Sea that allow for the growth of seagrasses. With the global decline in seagrass populations due to anthropogenic activity (Orth, Carruthers et al. 2006), research on the seagrasses of Red

Sea sharms provides a rare glimpse of a relatively pristine ecosystem, and offers an opportunity to survey the organisms coexisting with them. The biofilm of seagrasses has an abundance of epiphytic bacteria (Hemminga M. 2000). It is our suspicion that

Planctomycetes, extraordinary bacteria of the Planctomycetes-Verrucomicrobia-

Chlamydiae (PVC) superphylum, grow as part of the biofilm of seagrasses. The resulting thesis work is a study of three different seagrasses, Cymodocea serrulata, Halodule uninervis, and Halophila stipulacea, sampled from Rabigh Harbor and a coastal lagoon named Al Kharrar, and the subsequent isolation and purification of Planctomycetes bacteria from their biofilms for further molecular and computational work.

A word on lagoon and harbor ecosystems 14

Lagoon ecosystems are dynamic sites with complex nonlinear biological, chemical, and hydrological interactions. Parameters such as growth of primary producers, organic detritus load, nutrient recycling, and abiotic factors such as tidal flow rate, light intensity, wind speed, and salinity all serve as constraints for the functional health of the ecosystem. A shift in any one of these variables, or introduction of anthropogenic activity, can trigger massive changes in the state of the lagoon ecosystem, manifested by changes in species composition (Cioffi 2008). Their susceptibility to these sudden regime shifts makes them a fascinating environment to research.

Harbors, in contrast, are by definition places where ships, barges, and boats can dock to take refuge from wind, waves, and currents (Jellett 2012). They can be artificially constructed or natural, and may or may not contain a port or facility for loading or unloading vessels. Because the presence of sea vessel traffic is inherent in the definition, the ecosystem(s) contained within the harbor can be assumed to have been influenced to a greater extent by anthropogenic activity than lagoons, especially when they contain ports.

The two locations of our three collection sites were Al Kharrar Lagoon and Rabigh

Harbor, which contains Rabigh Industrial Port. To date there have been no microbiological or molecular ecological studies of either location, and data on hydrological circulation patterns, nutrient cycling, and eutrophication levels is limited or nonexistent. Nevertheless, the following descriptions should be useful in painting broad 15 strokes in order to visualize the settings from which the research materials were gathered.

Al Kharrar Lagoon

Al Kharrar lagoon, the collection site for one of our samples, is located between 22°45ʹ and 23°00ʹN latitude and 38°45ʹ and 39°00ʹE longitude along the western coast of Saudi

Arabia (Hariri 2008)(Fig. 1). To the south and east lie inter- and supratidal flats, known locally as sabkhas (Abd and Awad 1991). A narrow channel along the northwestern side extends for 20 km, parallel to the coast1. Red Sea sharms are believed to have been created by Late Pleistocene erosion followed by flooding due to rising post-glacial sea levels (Washmi 1999). Wadis, or seasonal streams in colloquial Arabic, formed networks that even today run to the coastal lagoons. The sabkha extends for 3 km to the east, with subsurface seawater extending for 1 km. From the western side, waves break on the Al Kassara reef and then calmly enter the lagoon (Ouf 1994).

1 It should be noted that the Saudi coast guard modified the lagoon and artificially constructed the channel, according to King Abdullah University of Science and Technology’s Coastal and Marine Resources Core Lab. 16

Figure 1. Location of Al Kharrar Lagoon2 (Batang 2012)

Mangroves of the species Avicennia marina are scattered throughout the sharm. Their presence both reduces wave energy and serves to relocate sediment from fluvial wadi streams to the lagoon while also enriching the sediment with biogenic carbon (Basaham

2008). High velocity tidal currents can enter Al Kharrar at speeds up to 1 m-s-1, but tidal range is low, only 20-30 cm at most (Hariri 2008). In normal dry weather conditions from March through October, cooler Red Sea water of lower salinity flows into the

2 According to KAUST Coastal and Marine Resources Core Lab, the only available base map of the collection sites are of low resolution, only for use in large-scale mapping, and cannot resolve details along the marginal areas of the Red Sea. The lagoon and bay are deformed and the relevant features are not resolved at the desired scale, i.e. the openings are not shown, shallow areas are not well-represented, outcrops (reefs and islets) are missing and unresolved, etc. As such, the Google Earth images (right of Fig. 1 and Fig. 5 in Methodology) are preferred as they clearly show the desired features and are the standard for scientific purposes until a better quality base map is made available. 17 sharm, and during the ebb tide, warmer water of higher salinity flows out (Basaham

2008).

Rabigh Harbor

To fully understand the context of the collection environment, Rabigh Harbor and its recent development have to be taken into consideration. Separated from Al Kharrar

Lagoon by approximately 11 km overland to the southeast, Rabigh Harbor is situated at the northern perimeter of Petro Rabigh, the refining and petrochemical complex. In addition to taking feed of 400,000 barrels of crude oil a day (PetroRabigh 2010), the complex contains a 1.3 million tonne/year ethane cracker, a 700,000 tonne/year monoethylene glycol plant, a 600,000 tonne/year linear low-density polyethylene plant, a 300,000 tonne/year high density polyethylene plant, and a 700,000 tonne/year polypropylene plant (Ling 2011). At the time of this work, the company had recently announced plans to double the petrochemical complex’s capacity over the next three years (Ling 2011).

With such a high level of anthropogenic activity introducing pollutants to the harbor the complex is situated upon, neither Rabigh Harbor nor Al Kharrar Lagoon can be referred to as pristine (Basaham 2008), as they once were in publications made prior to construction of the petrochemical complex in 2006. This makes an ecological survey of the seagrass species directly impacted by this activity an important contribution to marine science, because it may provide an early window to the decline in health of these populations over time. 18

Two out of three samples were collected from Rabigh Harbor, which is situated at

22°39’ N latitude and 38°59’E longitude (Fig. 2). It is 150 km north of Jeddah and only 30 km north of King Abdullah Economic City, a development project aiming to be the commercial hub of the kingdom. It is a deep-water port with an entrance channel 1300 m in length, 28 m in depth, and 240 m in width. There are four ports for tankers, small craft, yachts, and commercial cargo. The climate is that of a tropical desert, with humidity averaging 82% in the summer season, from March through October, and the average temperature ranging from 25° to 38°C. The majority of the time the prevailing winds originate from the North to Northwest with a speed of 2.6 – 12.9 m-s-1. The semi- diurnal tide cycle has a mean daily range of 0.34 m, with the port relatively unaffected by the general circulation of the Red Sea. Year round seawater temperatures range from 22° to 29° C (Aramco). 19

Figure 2. Location of Rabigh Harbor (Aramco)

Within the aforementioned environments of the lagoon and harbor grow economically and ecologically key populations of seagrasses.

Seagrasses: providers of crucial ecological services to marine environments

Seagrasses are flowering plants that have evolved over 100 million years to live fully submerged in the sea. They are crucial contributors to organic carbon production, nutrient cycling, and sediment stabilization (Larkum, Orth et al. 2006). They are believed to play an important role in trophic transfer to adjacent habitats and so are a 20 vital part of the food web where seagrass beds are located (Orth, Carruthers et al.

2006). From a phylogenetic perspective, seagrasses are unique for their low taxonomic diversity – only 60 species versus the 250,000 species of terrestrial angiosperms – yet high number of specific and profound adaptations that allow them to live in marine environments, including altered gas transport systems, chloroplasts, pollination mechanisms, and photosynthetic pathways. In spite of the fact that they have done a good job of gradually adapting over 100 million years to drastically shifting coastlines, varying sea composition and sea levels, and changes in atmospheric content, it remains to be seen whether they will be capable of keeping pace with the rapid contemporary changes exerted by human influence (Freeman, Short et al. 2008).

Seagrasses require up to 25% of incident light radiation to supply their roots and rhizomes with oxygen in the highly reducing environments in which they live, in comparison with <1% for terrestrial angiosperms (Dennison, Orth et al. 1993; Freeman,

Short et al. 2008). This requirement means that they are acutely responsive to environmental change, and are good indicators of water clarity. The United States government uses seagrasses as one of five sensitive bio-indicators of pollution (Orth,

Carruthers et al. 2006; Michael, Shin et al. 2008). Eutrophication and sediment runoff, introduction of invasive species, artificially induced hydrological changes, and commercial fishing have taken their toll on seagrass species, as reflected by the massive global decline in seagrass populations (Cambridge, Chiffings et al. 1986; Fourqurean,

Boyer et al. 2003; Marba, Santiago et al. 2006; Guimaraes, Cunha et al. 2012). Although 21 media coverage of the seagrass crisis has been lacking, with salt marshes, mangroves, and coral reefs receiving three- to 100 times the media attention, a fragmented but emerging network of seagrass ecologists is attempting to remedy the gap in public awareness by calling attention to these “coastal canaries” (Costanza, d'Arge et al. 1998;

Freeman, Short et al. 2008).

In many ways the health of the world’s coastal ecosystems depends on the services provided by the high value seagrass ecosystem. Seagrasses are an essential source of carbon, which is transported to the deep sea as a critical supply to food-limited environments (Duarte, Middelburg et al. 2005). Excess organic carbon produced is stored in the sediment and then sequestered in the biosphere (Larkum, Orth et al.

2006). Leaves, rhizomes, and roots determine the course of currents, trap nutrients, and filter input to coastal regions (Carr, D'Odorico et al. 2010). Species biodiversity in areas inhabited by seagrasses is recorded to be greater than areas with no vegetation

(Hemminga M. 2000). Faunal densities are found at a higher magnitude, and nursery grounds of many fish species are often located among seagrasses. Cross-habitat utilization with mangrove and reef ecosystems could be responsible for maintaining abundance of coral reef species (Beck, Heck et al. 2001). In most cases representative of coastal development, increased nutrition inputs (from wastewater or facilities such as petrochemical complexes) lead to large-scale loss of seagrasses, resulting in sediment re-suspension and thereby decreased fish populations (Orth, Carruthers et al. 2006). 22

Reviews of seagrass ecology and its methodology have criticized the descriptive nature of the majority of publications (Novak, Hines et al. 2009; Sghaier, Zakhama-Sraieb et al.

2011), calling for more complex syntheses and in-depth experiments to address the deficit in knowledge (Duarte 1999; Orth, Carruthers et al. 2006; Freeman, Short et al.

2008). The nutrient dynamics of seagrasses, and especially the effect of the epiphytes living in associated biofilms, is yet to be explored. It is known that seagrasses play an important role in nutrient reclamation, but the specific mechanism of nutrient resorption and the source of nutrient input – whether from leaf decay or microbial epiphytes – are not understood. There is evidence to suggest that parts of seagrass anatomy such as blades are hosts to aerobic nitrogen-fixing epiphytes, while the anaerobic roots are hosts to nitrogen-fixing anaerobic bacteria (McRoy, Goering et al.

1973; Larkum, Orth et al. 2006). Although a high amount of epiphytic photosynthetic and carbon-, nitrogen-, and phosphorus-transferring activity on the leaf surface of seagrasses has been observed (Michael, Shin et al. 2008), how the epiphytes contribute and the extent of their significance has yet to be elucidated. In tackling the seagrass knowledge deficit, determining the function of epiphytic bacteria is imperative if the scientific community is to gain a fuller picture of seagrass ecology.

A closer look at three marine angiosperm hosts

Cymodocea serrulata (Fig. 3) has linear leaves, 5-9 mm in width, which resemble straps.

The narrow base has a triangular leaf sheath, and the leaf tip is serrated. Scars are not 23 contiguous around the stem. They dwell in shallow, subtidal reef flats and sand banks

(McKenzie 2012).

Figure 3. C. serrulata as depicted by Seagrass-Watch (McKenzie 2012)

Halodule uninervis (Fig. 4) is from the same family as C. serrulata. It typically inhabits sublittoral areas but can live in intertidal areas. In sheltered, muddy environments it can have a broader leaf size of up to 3.5 mm, whereas in exposed, sandy shores the leaf size is as narrow as 1 mm. Due to its rapid colonization, it is very common (Environment

2007). 24

Figure 4. H. uninervis as documented by the Systematic Marine Biodiversity Inventory System (Environment 2007).

Halophila stipulacea (Fig. 5) come from a genus that lack basal leaf sheaths, unlike other seagrasses. They are able to adapt to a wide range of salinities. The rhizomes are branched and fleshy, and the roots are unbranched and thick with dense soft hairs.

Leaves are distributed in pairs along the rhizome, which takes root in sand, and range from 3 – 8 mm in width. Petioles can be 15 mm in length (Galil 2006). 25

Figure 5. H. stipulacea as identified by DAISIE Species Factsheet (Galil 2006)

In search of Planctomycetes: why seagrass?

As the focus of this work is to isolate Planctomycetes, it is important to expound the link between these prokaryotes and the eukaryotic system of seagrass biofilm within which we expect to find Planctomycetes. Seagrasses are living substrates for epiphytic colonization. The epiphytic community can be made up of autotrophs, including diatoms, cyanobacteria, and algae, and heterotrophs, including bacteria, fungi, and invertebrates (Corlett and Jones 2007). As nutrient enrichment is linked to increased epiphyte biomass, they are good bio-indicators of pollution. They are capable of matching and even doubling their seagrass host’s productivity and are involved in nutrient cycling. The spatially and biologically dynamic surfaces of seagrasses can provide epiphytes physiologically diverse microhabitats on the same leaf, and each host 26 can have differing surface chemistries that affect recruitment and distribution of colonies (Sirota and Hovel 2006). Seagrass epidermal cells are sites of nutrient and gas exchange, with invagination of their plasmalemmas possibly functioning in cell secretions. Up to two percent of photosynthetically fixed carbon is released as dissolved organic matter (DOM)(Moriarty, Iverson et al. 1986), and their release of nitrogen and phosphorous are likely to positively enhance epiphytic communities (Libes and Boudouresque 1987), making seagrasses beneficial hosts. As to whether or not epiphytes always benefit seagrasses, that matter is up for debate. Epiphytes potentially interfere in nutrient uptake by leaves and may act as competitors (Larkum, Orth et al.

2006).

Planctomycetes are not limited to epiphytic communities; they have also been found living as members of communities on, within, or in association with tiger prawn guts, oceanic abyssal sediments, marine snow, wild-gorilla feces, and in sewage (Fuerst,

Sambhi et al. 1991; Fuerst and Sagulenko 2011). Their habitats are remarkably wide- ranging as they can be found in freshwater and brackish water, and inhabit oligotrophic and eutrophic environments (Fuerst 1995). Relevant to this study are experiments in which Planctomycetes were found in association with macroalgae (Longford, Tujula et al. 2007; Lage and Bondoso 2011) and free-living as dominant lineages at times corresponding to diatom blooms (Morris, Longnecker et al. 2006). A species of the PVC superphylum, Pelagicoccus croceus, was isolated from seagrass (Yoon, Oku et al. 2007).

Finally, there are at least two publications linking Planctomycetes with seagrass 27

(Hempel, Blume et al. 2008; Ikenaga, Guevara et al. 2010). The recent work published on epiphytic community structure of marine plants and habitats of Planctomycetes strongly suggested to us that we would find Planctomycetes living on the biofilm of Red

Sea seagrasses.

The peculiarity of Planctomycetes, or why they are so special

Planctomycetes are a widely distributed but highly distinct group of bacteria belonging to the PVC superphylum. By the end of the last century, microbiologists were giving great attention to the unique morphological, structural, and genetic features that lent

Planctomycetes an aura of mystery and inspired much debate regarding their evolutionary history and physiological relevance. In brief, the following are some characteristics that set Planctomycetes apart from any other bacterial phylum:

Peptidoglycan-less: The cell walls of these bacteria lack a polypeptide chain that is found almost universally in other bacteria. Also known as murein, peptidoglycan strengthens most bacterial cells structurally and also plays a role in binary fission. The only other known bacteria to lack peptidoglycan are Chlamydiae and members of the Mollicutes class, including the genus Mycoplasma, which lack cell walls entirely anyway (Lindsay,

Webb et al. 2001). Instead, Planctomycetes have walls dominated by protein, with an abundance of cysteine and proline. Cysteine-rich outer membrane protein 2 (Omp2) is a protein homologous to one produced by chlamydiae and just happens to be a target in response to chlamydial infection (Strous, Pelletier et al. 2006). 28

Crateriform structures: One of the distinguishing morphological features of

Planctomycetes, the more so because it is easily seen using electron micrography, is the presence of indentations accompanied by raised rims, distributed either over the whole cell surface or just around the polar reproducing end, as is the case with Pirellula strains.

This trait has been used to justify the inclusion of morphotypes thought to be

Planctomycetes as true members (Fuerst 1995).

Budding division: Planctomycetes reproduce by budding (Fuerst 1995), not binary fission, with the exception of the proposed class Phycisphaerae (Fukunaga, Kurahashi et al. 2009) which has yet to be taxonomically accepted (Euzeby 2010). This is highly unusual for bacteria and something thought to be unique to yeast among single-celled organisms until recent decades. The mechanism by which this occurs is as yet unknown.

The phylum of Planctomycetes comprises one recognized class, that of Planctomycetia, which includes two orders, Planctomycetales and Brocadiales. Planctomycetales has one family thus far, the Planctomycetaceae, made up of nine recognized genera, the majority of them monospecific. They are Blastopirellula, Gemmata, Isosphaera,

Pirellula, Planctomyces, Rhodopirellula, Schlesneria, Singulisphaera, and Zavarzinella3.

Brocadiales has one family, the Brocadiaceae, containing “Candidatus” species (Garrity

2010). The special ammonium oxidizing properties of “Candidatus” Brocadia,

Anammoxoglobus, Jettenia, Kuenenia, and Scalindua species are discussed below.

3 Zavarzinella was described after the deadline for this source but Volume 4 of Bergey’s Manual still classifies it within the Planctomycetaceae family. 29

Even of the known species of Planctomyces, only a limited number have been studied using culture-dependent methods because most cannot be isolated in pure culture, and instead have been studied with culture-independent methods or using enrichment samples from the environment. Perhaps one reason for this failure to obtain isolates is the degree of metabolic diversity represented across species of Planctomyces. The media selected limits the type of metabolism that can survive from the start. The genera listed above can be categorized as mesophilic heterotrophic aerobes, moderate thermophilic, obligate oligotrophs, acidophilic heterotrophic aerobes, heterotrophic facultative anaerobes, and autotrophic anaerobes (Fuerst and Sagulenko 2011). Ward’s descriptions of the genera that scientists have managed to culture, from the family

Planctomycetaceae, range from ovoid to spherical to ellipsoid in shape; the cells can be single or in rosettes4; stalks can be absent, present, short or long; they can have or lack gliding motility; they can have single flagellums, flagella in polar bundles, or no flagella; they can have or lack pigmentation; and their salinity, pH, and temperature preferences can vary extensively. Marine, brackish, fresh-, ground-, and saltwater habitats; sewage, manure, compost, peat, and soil environments; as well as lice, termite, coral, sponge, prawn, and human colon tissue have all been sites where Planctomycetes have been observed and/or isolated (Garrity 2010). Given the diversity of habitats represented among such a short list of species, and a collectively poor understanding of the

4 In the case of Planctomycetes, a rosette means a multi-cellular organization where cells are joined together by non-cellular stalks. For a good example of this formation see Figure 3 of Kulichevskaya et al. (2012). Kulichevskaya, I. S., Y. M. Serkebaeva, et al. (2012). "Telmatocola sphagniphila gen. nov., sp. nov., a novel dendriform planctomycete from northern wetlands." Frontiers in Microbiology 3.

30 mechanisms of their metabolic pathways, it is no wonder that so few have been properly cultured and documented. Nevertheless, the most dramatic feature of

Planctomycetes, their cellular compartmentalization, has still been thoroughly described and has far-reaching ramifications for evolutionary theory.

Intracellular compartmentalization

Planctomycetes’ most striking characteristic is their intracellular membrane compartmentation. Although the extent to which the cytosol is compartmentalized can vary between species, they share a common pattern of cellular organization that calls into question the standing of the prokaryote-eukaryote dichotomy. The advent of light microscopy and its ability to detect nuclei launched an era when scientists categorized organisms as monera or eukarya based on their ability to detect organelles such as nuclei, mitochondria, and chloroplasts (Stanier and Niel 1962). The discovery of a shared cellular framework in Planctomycetes that includes a membrane-bound nucleoid blurs the lines between the three domains of life that biologists have referenced for the better part of the last century.

Thin sections of cryosubstituted and freeze-fractured cells have revealed all cultured and some uncultured species to be divided into different compartments by a major intracytoplasmic membrane (ICM), or a single bilayer within a Planctomycete’s cytoplasm separating the paryphoplasm from the regions of the cytoplasm which contain ribosomes (Lindsay, Webb et al. 2001; Fuerst 2005). It is completely distinct from the cytoplasmic membrane, and thin sectioning shows there is no contact between 31 the two membranes. A paryphoplasm, not to be confused with a periplasm, is defined as a region of the cytoplasm of a Planctomycete between the ICM and the cytoplasmic membrane which does not contain ribosomes (Fuerst and Sagulenko 2012). The compartmentalization is most complex in Gemmata obscuriglobus, where an additional double-membrane envelope, forming what has been termed the nuclear body, surrounds the genetic material of the cell. Immunogold labeling of DNA has shown that all Planctomycetes have condensed genetic material, an unusual trait for bacteria

(Lindsay, Webb et al. 2001), suggestive of proteins homologous to eukaryotes that function in organizing DNA. In the simplest compartmentalization scheme, representative of Isosphaera, Pirellula, Rhodopirellula, and Blastopirellula, the ICM divides the cytoplasm into an outer paryphoplasm and an inner pirellulosome. The pirellulosome contains the highly condensed nucleoid and ribosomes (Fuerst 2005).

Whether the paryphoplasm forms a polar cap region or the ICM invaginates to take up

60% of the cell depends on the genera, but all cultured Planctomycetes have a pirellulosome or riboplasm bound by an ICM, with a paryphoplasm between the ICM and cytoplasm (Fig. 3a and b)(Fuerst 2005). ANRV253-MI59-14 ARI 4 August 2005 12:34

32

Figure Figure 16. Diagram of the cellular organization of a) Pirellula and Isosphaera and b) “DiagramsCandidatus of cell” organization Brocadia anammoxidans and compartmentalization and in (aGemmata ) Pirellula (e.g.,from FuerstPirellula staleyi) and (2005) Isosphaera (e.g., Isosphaera pallida; plan also applies to Planctomyces maris) and (b)“Candidatus Brocadia anammoxidans,” and Gemmata (e.g., G. obscuriglobus). The varieties of cell compartmentalization found in different planctomycetes, as well as the underlying similarities in topology of their internal organization In the case of(i.e., possession of paryphoplasmthe “Candidatus compartment” species, an additional compartment is present for a total and intracytoplasmic membrane), are shown.

Annu. Rev. Microbiol. 2005.59:299-328. Downloaded from www.annualreviews.org of three compartments. The the ICM (Figure 1a). The paryphoplasmanammoxosome re- rest of the, believed to hold the enzymes required paryphoplasm surrounding the rest

by King Abdullah University of Science and Technology on 06/11/12. For personal use only. gion should not in any way be confused with a of the pirellulosome and its bounding single for anaerobic ammonium oxidationperiplasm, which is a layer between the cy- (Fuerst 2005membrane (48)), is bounded by a single membrane, (Figure 2). This structure ap- toplasmic membrane and cell wall, and the pears to be conserved in the closely related which is surrounded by cytoplasm containing ribosomes (termed the riboplasm) and detection of RNA within the paryphoplasm P. staleyi and B. marina and is similar (but with is consistent with its status as a true part of a slight variation) to the related Pirellula-like the cytoplasm of the planctomycete cell (48, planctomycete R. baltica. The phylogenetic fibrillar nucleoid. 50). The pirellulosomeEncompassing all of this is contains ribosome- significancethe ICM, which in turn is surrounded by the and predictive value of the struc- like particles and the cell nucleoid, which is ture are indicated by a study of a strain orig- paryphoplasmusually highly condensed. (Fig. 3b) In. The Pirellulaanammox staleyi inallyosome classifiedis an interesting case as a planctomycete on, not only mor- because and Blastopirellula marina (formerly Pirellula phological grounds, ATCC 35122, which was marina) the paryphoplasm forms a polar cap later confirmed as closely related to the type the enzymeregion at one pole hydroxylamine oxidoreductase that is continuous with the strain of(HAO) P. staleyiwas shown to be strictly confined ,ATCC 27377. Thin sections

306within its boundaries, but because it also has 13 Fuerst nm tubules, visible in some kind of 33 packaged arrangement (Lindsay, Webb et al. 2001). These observations have led those studying anammox to refer to the anammoxosome as a multifunctional compartment, possibly involved in cell division, chromosome replication, and other physiologically significant processes in addition to the ammonium oxidation process, to be described later (Jetten, Wagner et al. 2001).

Evidence of nuclear transport

Transmission electron micrography (TEM) of G. obscuriglobus showed that the membrane-bound nuclear body appeared to have clearly visible and significant gaps between the outer and inner membranes of the envelope. This topological clue implied that the apposed membranes might fuse to form a true folded single membrane, analogous to the eukaryotic nuclear envelope (Fuerst 2005). If this is the case, and the nuclear body is closed off from the cytoplasm and the ribosomes it contains by an envelope displaying continuity with itself as a single folded membrane, then there must be some way for RNA to reach the riboplasm from the nuclear body. In other words, the compartmentalization of G. obscuriglobus directed evolutionary biologists to look for evidence of membrane trafficking and energy-dependent endocytosis even though this process is thought to be unique to eukaryotes.

Recent studies have indeed demonstrated the ability of Planctomycetes to internalize proteins in an energy-dependent manner, compartmentalize and degrade proteins in the paryphoplasm, and associate the internalized proteins with vesicles (Lonhienne,

Sagulenko et al. 2010). The antibodies, used to react with the vesicle-like structures 34 that invaginated from the cytoplasmic membrane, bound to gp4978, a possible homolog of eukaryotic clathrin. This is consistent with findings of Planctomycete proteins possessing secondary and tertiary structural similarity to eukaryotic clathrin and coat protein families (Santarella-Mellwig, Franke et al. 2010). Recent phylogenetic analyses have dated the development of endocytosis and endomembrane systems to before the last eukaryotic common ancestor (LECA). These findings taken together have prompted the hypothesis that the last universal common ancestor (LUCA) used endocytosis as a form of heterotrophic nutrition, and Planctomycetes retained this system after LECA branched off. Alternatively, in a eukaryote-late scenario, Planctomycetes ancestors invented a basic membrane trafficking process which was laterally transferred to LECA

(Fuerst and Sagulenko 2012). Of course, this is up for debate. Another possibility is that after LUCA, many eubacteria, including Planctomycetes, branched off before archaebacteria split from the phylogenetic tree, and LECA developed an endomembrane system accompanied by the nuclear pore complex (NPC) (Cavalier-Smith 2006). This does not address Planctomycetes’ obtainment of NPC-like homologs (nup85 and nup133), which still others have suggested were passed through vertical descent from an ancestral Planctomycete to archaeal and eukaryotic counterparts (Devos 2012).

Because of the structural organization of the Planctomycete cell, some translation must take place in a non-DNA containing compartment, in a manner similar to eukaryotes.

The condensed nucleoid must be the site of transcription, and the mRNA must be transported through the nuclear envelope of the nuclear body. The presence of 35 membrane coat (MC) and nucleoporin homologs operating in endocytosis support this structure-based hypothesis, as does the ability of domains found in MC proteins to deform membranes, the presence of which is exclusive to the PVC superphylum among bacteria (Santarella-Mellwig, Franke et al. 2010). The ribosomes of Gemmata are bound in linear arrays on either side of the nuclear envelope membrane, suggesting that proteins produced via co-translation are secreted into the lumen of the nuclear envelope, analogous to the eukaryotic endoplasmic reticulum lumen (Fuerst and

Sagulenko 2012).

Also of significance to the debate over the evolution of endomembrane systems is the presence of sterols in Gemmata. Lanosterol and parkeol, its rarely found isomer, point to the retention of a primitive sterol pathway attained through lateral gene transfer

(LGT) from a unicellular eukaryote (Pearson, Budin et al. 2004). They could function to maintain internal membrane fluidity (Fuerst 2005), which for Planctomycetes would have been a characteristic critical for survival due to their compartmentalized cell plan.

Genome analyses

The origins of the endomembrane system are but one of many discussions sparked by unique Planctomycete traits. Genetic studies have uncovered a plethora of controversial data, some of it changing current perceptions of evolutionary origin, and some of it promising to revolutionize industry.

Archaea-like C1 transferase genes 36

The first two genomes to be completely sequenced were Rhodopirellula baltica, an aerobic marine Planctomycete, and aerobic freshwater G. obscuriglobus. Genome analyses were of particular interest because Planctomycetes' presence as significant members of far-ranging ecological communities begged the question as to their physiological potential. Annotation studies quickly revealed the surprising presence of tetrahydromethanopterin (H4MPT)-dependent enzymes (Glockner, Kube et al. 2003).

Prior to the discovery of these enzymes in aerobic methylotrophic proteobacteria

(Chistoserdova, Vorholt et al. 1998), it was thought that they only existed in the specialized energy C1 metabolism of anaerobic methanogenic archaea. The archaea-like enzymes Fae (formaldehyde-activating enzyme), Mch (methenyl-H4MPT cyclohydrolase), and Ftr/Fmd (formylmethanofuran:H4MPT- formyltranserase/formylmethanofuran dehydrogenase) are involved in a methylotrophic detoxifying pathway that oxidizes formaldehyde to formate and allows growth on C1 compounds (Marx, Chistoserdova et al. 2003). Because homologs of these archaeal proteins were only known to exist in proteobacteria, it was assumed that LGT from archaea was responsible, and that it had occurred in a single interdomain gene transfer event because of the clear separation between bacteria-specific genes and those with archaeal counterparts (Bauer, Lombardot et al. 2004).

When homologs were found in R. baltica, the matter of how archaea-like methanogenesis genes were transferred became a heated topic of discussion. Not only did the first two sequenced Planctomycetes genomes encode Fae and H4MPT- 37 dependent enzymes, but they also carried H4MPT/H4F-methylene dehydrogenase MtdA, highly similar to an enzyme from methylotrophic proteobacteria (Bauer, Lombardot et al. 2004). They also contained β-RFAP synthase (β-ribofuranosyl-aminobenzene-5'- phosphate) genes for catalyzing a methanopterin biosynthesis reaction, a co-factor upon which the activity of the other C1 transferases depends. In examining genome arrangement, it was found that while methanolotrophic proteobacteria had genes for the detoxifying formaldehyde oxidation pathway that were highly clustered and conserved, the counterparts of these archaea-like genes in Planctomycetes were scattered and the sequence of the gene modules was different (Bauer, Lombardot et al.

2004).

It remains to be seen whether or not these genes are even functional, and what their presence means in regards to Planctomycetes' metabolism. They cannot grow on common C1-substrates, and formaldehyde, which ought to be generated as an intermediate in the oxidation pathway, has not been detected (Chistoserdova, Jenkins et al. 2004; Woebken, Teeling et al. 2007). No known homologs for genes involved in the final steps of methanogenesis or C1 compound assimilation have been found in

Planctomycetes genomes (Bauer, Lombardot et al. 2004), thus genome analysis predicts that Planctomycetes by themselves are not capable of either methylotrophy or methanogenesis. Using normalized codon usage analysis, however, it has been shown that the fae1 gene is one of the most highly expressed of all Planctomycetes genes

(Bauer, Lombardot et al. 2004; Woebken, Teeling et al. 2007). Even in the absence of C1 38 substrates, all seven H4MPT-linked C1 metabolism genes are expressed in R. baltica.

Genome analysis may therefore be unable to recognize alternative cascades of the pathway that would be better discovered through experimental testing.

The question remains as to how these genes, traditionally viewed as components of an

"archaeal genomic signature," ended up in this phylum. An archaeal prototype from which such a transfer might have occurred has not been found (Chistoserdova, Jenkins et al. 2004). Cavalier-Smith (2006) has postulated that an LGT event between a proteobacterial methylotroph and a euryarchaeon took place, but this suggests that methylotrophy pre-dated anaerobic methanogenesis. This is contrary to the understanding of pre-historic Earth's atmosphere that most adhere to. Based on the topology of phylogenetic trees constructed using mch, chosen because of its essential function in C1 transfer, its lack of duplication in known genomes, and a lack of substitution by other enzymes, five alternate scenarios elucidating the relationship between Euryarchaeota, Planctomycetales, and Proteobacteriales have been put forth

(Chistoserdova, Jenkins et al. 2004).

1) The genes were transferred from Euryarchaeota to Bacteria through LGT before

Planctomycetes and Proteobacteria separated.

2) The genes transferred through independent LGT events after the separation of

Planctomycetes and Proteobacteria. 39

3) The genes originated in Proteobacteria and were independently transferred to

Euryarchaea and Planctomycetes (similar to Cavalier-Smith's hypothesis).

4) The genes were transferred independently from Planctomycetes to an early Archaeon and an early Proteobacteria.

5) The genes were present in LUCA before the branching off of Archaea from Bacteria, and most Bacterial phyla experienced gene loss.

Phylogenetic analysis is most in agreement with scenarios four and five because it matches the topology of mch-based trees and is consistent with the clear separation of archaeal and proteobacterial sequences. Because Planctomycete and proteobacterial sequences show both a clear distinction and with a degree of order conservation, and because methanogenesis could not have arisen prior to methanotrophy, scenarios one, two, and three are less likely (Chistoserdova, Jenkins et al. 2004).

Bearing in mind the antiquity of the C1 transfer pathway, if the origin of the methanopterin/methanofuran-linked pathway lies with Planctomycetes as scenarios four and five indicate, then Planctomycetes has a position near the root of the universal phylogenetic tree. Many groups studying this unique phylum have reached the same conclusion (Brochier and Philippe 2002; Chistoserdova, Jenkins et al. 2004) because

Planctomycetes seem to have a more ancient version of the C1 transferase genes than

Proteobacteria (Bauer, Lombardot et al. 2004), but a definitive conclusion is nowhere in sight. Bauer et al. (2004) proposes that an Archaeon transferred the genes through LGT 40 to a bacterial ancestor before the divergence of Planctomycetes and Proteobacteria. No one scenario is unequivocally the best.

Sulfatase activity

As early as 2002, Planctomycete genomic libraries were being searched using sequence tags in hopes of discovering genes that would provide clues as to their metabolic, physiological, and ecological significance (Jenkins, Kedar et al. 2002). They were the beginnings of the hunt to find homologous genes between the domains of life, even though the extent to which Planctomycetes genomes are littered with anomalies was yet unknown. By 2007 a multitude of sulfatases were discovered in a cross-comparison analysis using two metagenome libraries and sequencing from fosmids containing 16S rRNA genes (Woebken, Teeling et al. 2007). The high number of sulfatases found in

Planctomycetes may denote a part of their ecological role that has long been overlooked.

In freshwater Planctomycetes, such as Gemmata, there are only a few sulfatases, but in marine species such as Planctomyces maris, Blastopirellula marina, and R. baltica, there is a surprisingly large number. R. baltica has 109 sulfatases, more than any examined bacterial genome. When the Planctomycete-like fosmid sequences from this study were analyzed, taken together they carried genes encoding an average of 22 sulfatases per

Mb, a density even greater than that of R. baltica's genome (Woebken, Teeling et al.

2007). 41

The implications may well elucidate a potential role for marine Planctomycetes.

Sulfated heteropolysaccharides are abundant in marine environments (Fuchsman,

Staley et al. 2012). They are a component of chondritin in fish cartilage and are produced by red and brown algae (Araujo, Oliveira et al. 2004). Wallner et al. (2005) showed that R. baltica has proven sulfatase activity. Although R. baltica cannot degrade agar, it does anabolize carrageenan, another compound found in red algae (Woebken,

Teeling et al. 2007). The evidence suggests that sulfatase-rich Planctomycetes are well- suited for life in nutrient rich environments such as marine upwelling systems, as well as oxygen-limiting conditions. An increasingly popular belief is that Planctomycetes break down sulfated heteropolysaccharides and utilize their carbon skeletons as a food source

(Glockner, Kube et al. 2003; Woebken, Teeling et al. 2007; Fuchsman, Staley et al. 2012).

In contrast, anammox Planctomycetes, unlike members of the Planctomycetacia class, have a very different metabolic pathway.

Anaerobic ammonium oxidation

In the early 1990s, interesting observations were made at a denitrifying pilot plant - ammonium was disappearing along with nitrate, while nitrogen production was increasing. At the time, microbiologists thought it was impossible for the oxidation of ammonia to take place under anything but oxic conditions, but they soon learned that anaerobic oxidation is actually a much more favorable reaction (Jetten, Wagner et al.

2001). 42

First attempts to isolate the bacteria responsible for this process, termed "anammox"

(short for anaerobic ammonium oxidation), began by enrichment on mineral medium with ammonium, nitrite, and bicarbonate as the only carbon sources. All classical isolation methods failed, however, and in lieu of obtaining pure culture, microbiologists settled for creating a density gradient through centrifugation to achieve a sample with one contaminating bacteria for every 200 - 800 anammox bacteria (Jetten, Wagner et al.

2001). These were named “Candidatus” Brocadia anammoxidans, after the Gist

Brocades denitrifying plant where this process was accidentally discovered (Kuenen

2008). Morphological and genetic analysis soon identified the species as a deep- branching member of Planctomycetes.

After 15N-labeling experiments revealed that the oxidation of ammonium was very different from the aerobic oxidation process, it was proposed that hydroxylamine, a compound related to ammonium, reacted with ammonium to produce the intermediate hydrazine (Jetten, Wagner et al. 2001). The presence of hydrazine, an energy-rich compound that can be used for rocket fuel, was indeed obtained in enrichment cultures

(Kuenen 2008).

Those studying anammox bacterium soon encountered setbacks. The doubling time of

“Candidatus” B. anammoxidans was three weeks, and even by the early 2000s with a better understanding of the organism's metabolism the doubling time was only reduced to two weeks (Jetten, Sliekers et al. 2003). Today it is possible to bring their doubling time down to eight days, but only in suspension under highly enriched conditions 43

(Kuenen 2008). Still, “Candidatus” B. anammoxidans oligonucleotide-specific probes were developed for fluorescence in situ hybridization (FISH) and tested in wastewater systems. Surprisingly, the probes bound to bacteria participating in anammox reactions, but none of the species identified were “Candidatus” B. anammoxidans. A new genus,

“Candidatus” Kuenenia stuttgartiensis, was discovered, with 90% rDNA similarity to

“Candidatus” B. anammoxidans (Jetten, Wagner et al. 2001). This was soon followed by discovery of a third anammox species, “Candidatus” Scalindua sorokinii, in the Black Sea, the first species found in nature to anaerobically remove fixed inorganic nitrogen

(Jetten, Sliekers et al. 2003).

Genome analysis revealed biochemical and physiological properties that were later confirmed by enzyme assays. It was shown that acetyl-CoA is involved in the fixation of carbon dioxide, such that one molecule is reduced to a methyl group and another is reduced to carbon monoxide; they then combine to yield acetyl-CoA, which is further reduced to pyruvate. The oxidation of hydrazine provides the low redox potential electrons to fuel this high energy reaction (Kuenen 2008). A putative gene for a nitric oxide-producing nitrite reductase was discovered, as well as a gene encoding HAO, giving rise to current understanding of the oxidation mechanism whereby these enzymes form hydroxylamine from nitrite or nitric oxide. Nitric oxide and ammonium combine to form hydrazine, which reduces ferredoxin so that a proton motive force

(PMF) is generated across the previously discussed anammoxosome membrane (Fig.

4a). PERSP ECTIVES 44

a b At the aerobic–anaerobic interface – + – – + (for example, in a biofilm, in a sediment NO2 NH4 2NO3 3NO2 NH4 or in stratified water bodies), interest- ing interactions and competition can NIR NO HH N2H4 HAO N NAR NIR NO HH N H 2 2 4 occur between the anaerobic anammox 1e– 3e– 4e– 1e– 3e– bacteria and the aerobic ammonium- and nitrite-oxidizing bacteria — the aerobic

– nitrite oxidizers compete with the aerobic 4e N 2 ammonium oxidizers for oxygen, and 6H+ 3H+ 8H+ 8H+ 6H+ the anammox bacteria compete with the + + + + + + ammonium oxidizers and nitrite oxidizers Anammoxosome bc Q ATPase NAR NAR Q bc HD 27 membrane 1 1 for ammonium and nitrite, respectively – – – – – – (FIG. 5). Notably, the anammox bacteria ATP 4e– and aerobic nitrite-oxidizing bacteria at this interface require aerobic ammonium- Ferredoxin oxidizing bacteria to produce one of their Figure Figure 73. Schematic of possible anammoxosome membrane reactions involved in | Hypothetical catabolism and reversed electron transport in the anammoxosome. substrates: nitrite. Experiments that used catabolism and a | Pathway of ammoniumreverse electron transport chain oxidation that uses nitrite as the electron acceptor for the creation of an oxygen-limited reactor, which was fed a proton motive force (PMF) over the anammoxosomal membrane. NaNitriteture Re (NOviews–) is| Micr reducedobiolog toy a) Nitrite (NO2-) is reduced to nitric oxide (NO), which combines with ammonium (NH2 4+) with ammonium only, revealed that at low to yield hydrazine (Nnitric oxide, which thenH combines). This requires one and three electrons, respectively. When with ammonium to produce hydrazine, with the uptake of one oxygen concentrations (up to 2 mg per plus three low-energy2 electrons.4 The oxidation of hydrazine to nitrogen yields four high-energy hydrazine is oxidized to nitrogen, four electrons are produced and enter the quinone electrons, which flow downhill through the quinone (Q) pool and the H+-translocating cyto- litre in the bulk phase) coexisting small pool (Q) before being transferred to the cytochrome bc1 complex. This serves to clumps of Candidatus B. anammoxidans chrome bc1 complex, thereby generating a PMF that is inside positive. The PMF energizes the generate a PMF that gives Aproton-translocating ATPase forTPase the energy to p the production of ATProduce ATP in the riboplasm. b) in the riboplasm. The electrons are recy- and separate clumps of a Nitrosomonas eutropha strain dominated the reactor Reversed electron transport powered by PMF showing how nitrate reductase (NAR) cled from quinone–cytochrome bc1 oxidoreductase the hydrazine-forming reactions. b | PMF-driven – biomass28,29. All of the removed ammonia oxidizes nitrite and yields electrons that are reversed electron transport combines central catabolismthen donated back with nitrateto the anammox (NO3 ) reductase to gener- ate ferredoxin for carbon dioxide reduction in the acetyl-CoA pathway. Hydrazine can donate reaction in order to generate reduced ferredoxin that can participate in the acetyl-CoA was converted to N2 gas by a coupled nitrite- high-energy electrons to ferredoxin, but these electrons are not recycled. Nitrite oxidation to pathway. HAO = hydrazine oxidoreductase; HD = hydrazine dehydrogenase; HH = formation–anammox reaction, and only a nitrate through nitrate reductase (NAR) compensates for this, but yields low-energy electrons few nitrite oxidizers could be detected. This hydrazine hydrolase; NIR = nitrite oxidoreductase that must be ‘energized’ by the PMF to be fed back into(Kuenen 2008 the anoxic ammonium) oxidation (anam- oxygen-limited combined process for the mox) reaction. This ‘energization’ is accomplished by a nitrate reductase that operates at the removal of ammonium has been patented expense of the PMF. HAO, hydrazine oxidoreductase; HD, hydrazine dehydrogenase; HH, hydra- (the CANON (completely autotrophic Alternatively, ferredoxin can donate electrons to the acetyl-CoA pathway for carbon REF. 18 zine hydrolase; NIR, nitrite oxidoreductase. Figure modified, with permission, from Nature 30,31 ‘ (2006) Macmillan Publishers Ltd. nitrogen removal over nitrite) process) , dioxide fixation. Electrons donated from the nitrite-nitrate oxidation reaction replace and is discussed below. When the oxygen concentration was gradually increased, coexisting populations of three organisms, electrons lost in this process, but first they must enter the quinone pool via PMF-driven environments, such as the Black Sea, the capacity of Candidatus A. propionicus for including a nitrite oxidizer, were obtained. coasts of Namibia, Chile, Peru24,25, and, propionate metabolism gave it a competi- reversed electron transport (Fig. 4b). The bacterium deviates from the typical more recently, Lake Tanganyika, a fresh- tive advantage, and pointed to a clearly Environmental activity and global impact water lake in western Tanzania26. There defined ecological niche for this organism. In the early 1990s, the Gist Brocades denitrification reaction so drastically for this reason: both nitrate and nitrite can accept is also evidence of organisms that belong As mentioned above, the habitat of Fermentation Company closed their pilot to the Candidatus Scalindua family in anammox bacteria requires the simul- plant, leaving the reactor in our laboratory electrons to eventually form ammonium, and nitrate sediments off the coast of Gothenburg and taneouscan be converted into both nitrite presence of ammonium and as the only confirmed source of anammox the San Pedro basin (W. Berelson, unpub- nitrite, which can be found at or near the bacteria. Consequently, the Delft-Nijmegen and ammoniumlished communication). (Guven, Dapena et al. 2005 Recently, the ). This means that aerobic–anaerobicanammox interface can create its of sediments team13 and a Swiss team32 began to look for Jetten team successfully enriched for these and water bodies. Ammonium is produced evidence of the anammox reaction elsewhere, organisms from the Gullmarsfjord coastal by the anaerobic degradation of organic particularly in waste-water-treatment plants. sediment, Sweden, using a sequencing matter, both through ammonification The organisms were eventually detected fed-batch reactor. Brocadia and Kuenenia and dissimilatory nitrate and/or nitrite and identified in many waste-water-treatment species are the most commonly found reduction. The nitrite may originate from plants and a large range of marine and organisms in the enrichments from waste- nitrate reduction, which, in turn, may fresh-water habitats. Using 15N-nitrogen water-treatment plants and large-scale be due either to common denitrifying compounds, Thamdrup and Dalsgaard33 anammox reactors. Interestingly, a mem- organisms (lithoautotrophic or organo- looked for anammox activity in marine ber of a new anammox family, Candidatus heterotrophic) or nitrate reduction by sediments from Danish coastal regions, Anammoxoglobus propionicus, was the anammox bacteria in the presence using labelled ammonium or nitrate in an discovered in fed-batch enrichments (in a of organic compounds such as formate, isotope-pairing technique34,35. This analysis minerals medium that contained ammo- acetate or propionate. When ammonium provided the first clear evidence that anam- nium and nitrite) that were supplemented diffuses upwards and meets the oxygen mox activity was detectable in different with increasing concentrations of propion- that is diffusing downwards, the nitrite natural habitats; however, the convincing 21 ate . Competition experiments between could also be derived from nitrification by proof that the mixed-labelled N2 gas was this bacterium and Candidatus B. anam- aerobic bacterial or crenarchaeal ammonium due to anammox bacteria came from a cold moxidans demonstrated that the higher oxidizers. Christmas expedition to the Black Sea,

NATURE REVIEWS | MICROBIOLOGY VOLUME 6 | APRIL 2008 | 323 45 own inorganic electron acceptor as well as donor from various carbon sources. Through this process, nitrogen gas is released (Kartal, Kuypers et al. 2007).

After immunogold labeling revealed HAO to be confined within the boundaries of the anammoxosome, roughly 30% of the cell volume, further studies took a closer look at the composition of the specialized compartment's membrane. Unique lipids, termed ladderane lipids for their cis-ring junctions that form a staircase-shaped conformation of butane rings fused in a linear fashion, appear in all groups of anammox bacteria (Jetten,

Sliekers et al. 2003). Some of them are variations of methyl esters of a C20 fatty acid chain with five concatenated cyclobutanes; others are a C20 chain with only three cyclobutanes, a cyclohexane, and an sn2-glycerol monoether; yet others are mixed ether-ester lipids containing a combination of both ladderanes (Damste, Strous et al.

2002). Ether-linkages in addition to the ladderane moities were also detected, an unusual find because they were thought to be special to Archaea. The presence of mixed ether-ester lipids is one reason some have marked Planctomycetes as an early- diverging bacteria (Brochier and Philippe 2002).

The structure of the ladderane lipids sheds light on the complex biochemical and physiological reactions happening across the anammoxosome's membrane. It is a highly sterically strained compound rarely seen in nature, and seems specially adapted to allow anammox its unique metabolism. The unusually dense conformation of carbon atoms can form a tight barrier against diffusion, allowing toxic reagents like hydrazine and hydroxylamine to remain sequestered away from the rest of the cell, while 46 simultaneously maintaining a concentration gradient (Jetten, Sliekers et al. 2003). Since their discovery they have been used as signature molecules to detect anammox presence in marine environments (Kuenen 2008).

In nature

Anammox has been detected using 16S rRNA, FISH, ladderane composition analysis, and mixed-labeled nitrogen in the Black Sea, Costa Rica, off the coasts in upwelling zones of

Peru, Chile, and Namibia, and even in freshwater systems such as marshes, lakes, and peat bogs (Kuenen 2008). Some estimate anammox to be responsible for more than

50% of nitrogen removal from marine ecosystems (van de Vossenberg, Rattray et al.

2008). The importance of the ecological contributions made by “Candidatus” species cannot be overestimated. In continental shelf sediment ecosystems, up to 67% of the nitrogen gas produced is attributed to anammox bacteria (Kuenen 2008).

In industry

It was estimated within a few years of the discovery of anammox bacteria that operational costs of wastewater systems could experience a 90% reduction by saving energy and resources in removal of nitrogen (Jetten, Wagner et al. 2001). The prediction of stable and high quality effluent without the need for process control and the promise of compact ammonium removal was tempting, and after two years of preparation due to the slow doubling time of anammox bacteria, the first plant was fully operational. It exceeded expectations by performing at twice the capacity for which it 47 had been designed (Kuenen 2008). Several anammox reactors are currently in operation with more underway.

Research hypotheses and objective

Although they are nearly ubiquitous in nature, Planctomycetes are still as enigmatic as when they were first discovered and confused for fungi (Fuerst 1995). It seems the more the scientific community learns about this phylum, the more questions arise.

These organisms’ ability to anaerobically oxidize ammonium, their high level of sulfatase activity, and their participation in the global carbon cycle has launched many an evolutionary debate. Then there is the matter of their intracellular compartmentalization - how and when did this come about? The implications are limitless, not only for evolution but for industry as well. Until their ecological roles are better understood, however, microbiologists will arrive at an impasse regarding application. Intensive molecular, genetic, and bio-computational work remains to be done, but it all begins with isolation and cultivation.

The objective of this study was to isolate and purify Planctomycetes from biofilm samples collected from the Rabigh Harbor and lagoon seagrasses of the Red Sea.

We hypothesize that 1) Planctomycetes are present in the Red Sea and inhabit the microhabitats comprising the biofilm of seagrasses; 2) their isolation is possible using conventional enrichment and isolation methodology specially adapted for Red Sea microbes; and 3) through repeated colony picking and streaking we can obtain pure 48 culture of Planctomycetes for further studies, and this can be confirmed through morphological characterization and electron microscopy.

Achievement of this objective will add to the body of knowledge on Planctomycetes as this phylum has yet to be isolated from the Red Sea, and the strains that have so far been isolated in pure culture worldwide are few in number.

49

CHAPTER 2: METHODOLOGY

Media preparation

Enrichment broth

Selective broths were prepared in order to create aqueous environments advantageous for Planctomycetes and inhibitive of competing bacteria. Because of the absence of peptidoglycan in Planctomycetes' cell walls, the beta-lactam antibiotics streptomycin and ampicillin were chosen as selective agents against which Planctomycetes has demonstrated resistance (Fuerst, Gwilliam et al. 1997). Because of the slow doubling time of Planctomycetes, this was especially helpful because even as the antibiotics lost their effectiveness over the course of a week, the fast-growing, non-resistant competitors (the majority of gram-negative bacteria) of Planctomycetes were prevented from taking over plates.

M13 media

M13 media was prepared as follows, modified from Schlesner (Schlesner 1986; 1994), because it has been used consistently by microbiologists to successfully isolate

Planctomycetes (Gade, Schlesner et al. 2004).

Per liter:

Carbon sources:

0.2 g peptone from casein

0.2 g yeast extract

0.2 g glucose 50

Solutions:

20 ml Hutner's basal salts solution

10 ml Vitamin solution No. 6

50 ml 0.1 M Tris/HCl, pH 7.5

250 ml sterilized Red Sea water (SRSW)

Selective reagents:

1 g streptomycin from prepared stock solution

0.2 g ampicillin from prepared stock solution

40 mg cycloheximide from prepared stock solution

Double distilled water (dd H2O) was added up to one L (approximately 648 ml)

Hutner's basal salts solution:

10.0 g Nitrilo triacetate (NTA)

29.70 g MgSO4*7H2O

3.34 g CaCl2*2H2O

12.67 g NaMoO4*2H2O

99.0 mg FeSO4*7H2O

50 ml Metal salts "44" solution

900 ml dd H2O for a total volume of 1 L

NTA was prepared by dissolving ten g in roughly 300 ml dd H2O. It was then neutralized with 7.3 g KOH salts so that the precipitate disappeared. The pH was adjusted to 7.2 using 1 M KOH solution.

The solution was stored at 4°C. 51

Metal salts "44" solution:

250.0 mg ethylenediaminetetraacetic acid (EDTA) in free acid crystals, not salt

1,095 mg ZnSO4*7H2O

500.0 mg FeSO4*7H2O

154.0 mg MnSO4*H2O

39.2 mg CuSO4*5H2O

20.3 mg CoCl2*6H2O

Dd H2O was added up to one L and the solution was stored at 4°C. Drops of H2SO4 were added to retard precipitation.

Vitamin solution No. 6:

4.0 mg biotin

20.0 mg pyridoxine HCl

10.0 mg thiamine HCl

10.0 mg Ca-pantothenate

10.0 mg ρ-aminobenzoic acid

4.0 mg folic acid

10.0 mg riboflavin

10.0 mg nicotinic acid

0.2 mg vitamin B12

Dd H2O was added up to one L; the solution was sterilized by filtration only.

Tris: 52

One L of Tris was prepared by dissolving 12.114 g Tris in 500 ml dd H2O, adjusting the pH to 7.5 with a pH meter and 1 M HCl, and adding dd H2O up to one L.

SRSW was prepared by autoclaving two L glass containers with screw top lids at 121°C for 15 minutes.

Antibiotic stock solutions:

A 1 g/1 L final concentration stock solution of streptomycin was needed. It was calculated that for all enrichment and solid media experiments 20 L M13 solution would be required, so 4,000 mg streptomycin was dissolved in 40 ml dd H2O and filter sterilized.

A 0.2 g/1 L stock solution of ampicillin was needed for 20 L M13 solution, so 20,000 mg ampicillin was dissolved in 400 ml dd H2O and filter sterilized.

Both solutions were stored at 4°C.

Per liter of M13 broth prepared, two ml streptomycin and 20 ml ampicillin stock solutions were added to solution. See below.

Antifungal agent:

When adding 40 mg/L cycloheximide to each liter of M13 media, the two ml streptomycin and 20 ml ampicillin were first mixed in a beaker and the cycloheximide powder was dissolved as completely as possible in this solution before addition to the rest of the broth.

Difco Marine Broth 2216 plus antibiotics 53

Difco Marine Broth 2216 plus antibiotics (referred to as “anti-2216” for brevity) was utilized because it is nutrient rich compared to M13 media but still selective because of the beta-lactam antibiotics, so it made for good comparison. It was prepared by mixing

500 ml dd H2O with 37.4 g Difco Marine Broth 2216 powder, and adding dd H2O up to

978 ml. Using a magnetic stir bar and heating block at 180°C, the solution was heated until clear, usually one to two hours. The stir bar was removed with a sterile magnetic stir rod, and the beaker was autoclaved for 15 minutes at 121°C. When the solution cooled to 55°C, while it was still warm enough to pour but cool enough to touch, two ml streptomycin and 20 ml ampicillin stock solutions were added along with 40 mg cycloheximide, which was first dissolved in a beaker containing the antibiotics stock solutions to be added. Broth solution was kept at 4°C.

1/10 Difco Marine Broth 2216

1/10 Difco Marine Broth 2216 (referred to as “1/10-2216”) was used because instead of selecting for Planctomycetes using only antibiotics, like the anti-2216 broth, and instead of providing a nutrient-poor environment in addition to using antibiotics to select for

Planctomycetes, like the M13 broth, the diluted 2216 broth selected for Planctomycetes only on the basis of its nutrient-poor (oligotrophic) conditions. Studies have shown that

Planctomycetes can be selected for using dilute media (Staley 1973; Schlesner 1994). It was prepared by mixing 500 ml dd H2O with 3.74 g Difco Marine Broth 2216 powder, and adding dd H2O up to one L. Using a magnetic stir bar and heating block, the solution was heated at 180°C until clear, usually one to two hours. The stir bar was 54 removed with a sterile magnetic stir rod, and the beaker was autoclaved for 15 minutes at 121°C. Broth solution was kept at 4°C.

Solid media

Agar media

The above solutions were prepared as described, and in addition 18 g/L granulated agar was added and the solutions were heated on heating blocks at 180°C until the agar was completely dissolved, usually two hours, before being autoclaved at 121°C for 15 minutes. In the case of Difco Marine Broth 2216 plus antibiotics agar media, antibiotics and antifungal solution was not added until the media had cooled to 55°C, just prior to pouring the solution into plates. In the case of M13 agar media, Hutner's basal salts solution, Vitamin solution No. 6, and antibiotics and antifungal solution were mixed together and added to the media after it was autoclaved, once it had cooled to 55°C.

Gellan media

In addition to preparing M13 agar plates, M13 gellan gum plates were prepared as well.

The rationale behind this was that several studies have shown gellan gum to be more successful in isolating bacterial species that are difficult to culture (Davis, Joseph et al.

2005). Gellan is also clearer and freer of artifacts, desirable traits to have during the culturing process. Steps were the same for agar media preparation except in lieu of 18 g/L granulated agar, six g/L gellan gum and one g/L CaCl2*6H2O (to assist in dissolving) were added. Like the M13 agar media, for M13 gellan media the Hutner's basal salts 55 solutions, Vitamin solution No. 6, and antibiotics and antifungal solution were added after autoclaving at 55°C or cooler before pouring.

Full-strength Difco Marine Broth 2216 agar

Difco Marine Broth 2216 agar (full-strength; referred to as “2216”) was also prepared by heating 37.4 g/L Difco Marine Broth 2216 and 18 g/L granulated agar over a heating block set to 180°C until the agar had dissolved, and autoclaving at 121°C for 15 minutes.

Plate size

All media was poured into 100 x 15 mm polystyrene Petri dishes.

Collection of samples

Saudi regulations prohibited female members of the team from participating in the collection trips, but through correspondence with King Abdullah University of Science and Technology's (KAUST) Coast and Marine Resources Core Lab (CMOR), the following information was obtained (Batang 2012).

Rabigh Harbor

On April 9th and 10th, 2012, seagrass samples of species later identified as Cymodocea serrulata and Halodule uninervis were collected from the innermost, eastern section of the harbor (Fig. 5b, RB5) at depths ranging from two to four m. This section of the harbor contained a mixed-species seagrass bed, dominated by 40-70% coverage of C. 56 serrulata and adjacent to isolated patches or intermixed with H. uninervis with 20-30% coverage. H. uninervis inhabited deeper zones and tended to be less dense. Sparse patches of Halophila ovalis, roughly 5% coverage, also occurred on the offshore edges of the seagrass meadow. The meadow is adjoined to a channel lined with mangroves that leads further east to join a wadi. Due to this geography, the majority of seagrass blades were silt-coated and onsite substrate was fine-grained, soft, and highly bio-turbated with dense animal burrows.

Seagrass and sediment samples were obtained through assistance of Self-Contained

Underwater Breathing Apparatus (SCUBA). The seagrass samples consisted of entire shoots carefully cut off from the rhizomes. Samples were placed in sealed bags filled with onsite seawater to prevent abrasion of the shoots with the container. Sediment samples were collected from the same seagrass beds using pre-split polyvinyl chloride

(PVC) push cores, each 5 cm in diameter and 20-25 cm long, with removable caps at both ends. Cores were collected in triplicate. Seagrass and sediment samples were placed in sealed bags, and chilled in ice coolers until further processing at the laboratory on the same day. 57

Figure 8. Map of sampling sites at a) Al Kharrar Lagoon and b) Rabigh Harbor. Location and relative sizes are as shown in the inset. Satellite images provided by Google Maps; upper right map provided by KAUST CMOR (Batang 2012).

Al Kharrar Lagoon

This collection trip took place on April 11th, 2012. The site was densely vegetated with a single species of seagrass, Halophila stipulacea, from 5-8 m in depth, covering the central through the southern regions (Fig. 5, KL15). The monospecific beds comprised

80-100% of the coverage across the whole lagoon. The seagrass samples consisted of entire shoots carefully cut off from the rhizomes and were obtained through SCUBA.

Samples were placed in sealed bags filled with onsite seawater to prevent abrasion of the shoots with the container, and placed in ice-filled plastic coolers until further same-

5 Sampling site titles KL1 and RB5 are kept from the time of collection in order to be consistent with KAUST CMOR’s choice of labels, as samples for other experiments were also collected at this time. 58 day processing at the laboratory. The sediment samples were collected using PVC push cores in the same manner as Rabigh Harbor and kept on ice until processing at the laboratory.

Environmental Data

The following parameters were measured on site on the day of collection using a multiparameter sensor assembly Yellow Springs Instrument 6600 Sonde:

Table 1. Rabigh Harbor and Al Kharrar Lagoon Environmental Parameters Parameter Rabigh Harbor Al Kharrar Lagoon Temperature 26.93°C 27.51°C

Specific Conductivity 58.120 micro Siemens/cm 60.980 micro Siemens/cm

Conductivity 60.270 micro Siemens/cm 63.900 micro Siemens/cm

Total Dissolved Solids 37.780 TDS g/L 39.640 TDS g/L

Salinity 38.73 ppt 40.88 ppt

Depth 3.986 m 6.494 m pH 7.94 7.77

Oxygen Reduction Potential 103 mV 125 mV

Dissolved Oxygen 6.58 mg/L 6.07 mg/L

Coordinates 39 degrees 0' 35.762" E, 22 38 degrees 54' 39.638" E,

degrees 45' 5.582" N 59

22 degrees 54' 50.251" N

Processing of samples

Seagrass

As soon as the seagrass samples arrived from the collection trip team, collection of the biofilm began. Seagrass shoots were pressed against the rim of a 15 ml Falcon tube at approximately a 45° angle, in such a way as to avoid breaking the shoot by exerting too much strain. Razor blades were not used because only the epiphytic communities of the biofilm were the target, not endophytic communities, so care was given not to nick or cut the shoot. Instead, the shoot was dragged along the side of the Falcon tube so that the biofilm slid down and was collected as the shoot was pulled up. Care was taken to keep the plant intact. Between seven and ten ml biofilm solution was collected for each seagrass sample from both sites.

While processing the Halophila stipulacea samples, it was observed that some of the seagrass shoots were relatively normal, while others appeared to have unidentified white growths on them, roughly one cm in diameter. It was decided at this time to collect between seven and ten ml biofilm from the "normal"-looking seagrass shoots, and seven to ten ml biofilm from the shoots with the unidentified growths. The growths were easily picked off with a razor blade because they were not very strongly attached, and the biofilm was collected as described. From this point forward the two groups were treated as separate samples. 60

Enrichment

From each of the four categories of seagrass samples - C. serrulata, H. uninervis, H. stipulacea "normal," and H. stipulacea abnormal - one ml of the collected biofilm was placed directly into a 250 ml Erlenmeyer flask for enrichment in M13 broth, 2216 plus antibiotics broth, and 1/10 2216 diluted broth, for a total of 12 flasks. The flasks were placed on shaking incubators with a temperature of 28°C set at 220 rpm for the duration of the experiment.

Plating

Dilutions of the biofilm solutions were created by adding one ml of each biofilm sample to four ml SRSW in a 15 ml Falcon tube and mixing thoroughly. From this mixture, one ml was taken and added to four ml SRSW in a different 15 ml Falcon tube, thus creating

1:5 and 1:25 dilutions.

From each concentration (undiluted, 1:5 dilution, and 1:25 dilution ) of all categories of biofilm, 100 µl was spread evenly across each of the following plates: M13 agar, M13 gellan, anti-2216, 1/10 2216, and 2216. Samples were spread under a sterile fume hood using Becton Dickinson and Company disposable pre-sterilized L-shaped cell spreaders.

Plates were incubated at 28°C with the media side up to prevent condensation.

Sediment

There were three categories of sediment, one for each species of seagrass. From Rabigh

Harbor, the sediment associated with C. serrulata was medium brown with a soil-like 61 consistency. Sediment collected with H. uninervis was very dry and had inconsistent texture, chunky and crumbly in some parts while not in others. It was a darker brown color. From Al Kharrar Lagoon, the sediment collected with H. stipulacea was claylike, with finer granules, and grayish brown in color.

Ten g of each sediment sample was placed in a 50 ml Falcon tube, and ten ml SRSW added. After thorough mixing, one ml from each tube was added to four ml SRSW in a

15 ml Falcon tube and mixed to form a 1:5 dilution. Subsequently, one ml of that solution was mixed with an additional four ml SRSW in another 15 ml Falcon tube to form a 1:25 dilution, for a total of nine tubes.

Enrichment

One ml of each of the initial sediment plus SRSW mixture (i.e. undiluted) was placed directly into a 250 ml Erlenmeyer flask for enrichment in M13 broth, 2216 plus antibiotics broth, and 1/10 2216 diluted broth, for a total of nine flasks. The flasks were placed on shaking incubators with a temperature of 28°C set at 220 rpm.

Plating

From each concentration (undiluted, 1:5 dilution, and 1:25 dilution ) of all three categories of sediment, 100 µl was spread evenly across each of the following plates:

M13 agar, M13 gellan, anti-2216, 1/10 2216, and 2216. Samples were spread under a sterile fume hood using Becton Dickinson and Company disposable pre-sterilized L- 62 shaped cell spreaders. Plates were incubated at 28°C with the media side up to prevent condensation.

Isolation and Purification Techniques

Plates were observed and colony morphology documented on a rotating daily basis. If new and different colonies were noticed, they were picked with disposable pre- sterilized inoculating loops (Becton Dickinson and Company) and re-streaked using the four quadrant streak pattern onto a new plate of the same media. For each quadrant, a new sterile loop was used in order to increase the chances of isolating single colonies. If after a second or third passage isolation was not achieved, a subsequent passage was made, but usually isolation did not take more than three passages.

Enrichment broth to solid media

If interesting and/or cloudy growth, suspected to be bacteria and not fungal, was observed in the enrichment media during the first one to three weeks, 100 µl of the solution was pipetted from the broth and spread evenly across corresponding plates using pre-sterilized disposable L-shaped cell spreaders. For instance, bacterial growth from M13 broth was spread onto M13 agar and M13 gellan plates; growth from 2216 plus antibiotics broth was spread onto anti-2216 agar plates; growth from 1/10 2216 diluted broth was spread onto 1/10 2216 agar plates. In almost every case, in flasks with cloudy growth there was also sludge-like build-up on the side of the flask. This was collected with a sterile cotton swab and placed in 1 ml SRSW; 100 µl of this mixture was 63 then used to spread on corresponding plates. After growth on plates was observed, colonies were picked and re-passaged as described above.

Transmission Electron Micrography

After study under a light microscope of some specimens, it was suspected isolation of

Planctomycetes had been achieved but we wanted to confirm this with results from

TEM. In these cases, 2.5% glutaraldehyde solution was prepared using dd H2O and colonies were suspended in one ml of this solution in two ml screw cap tubes. They were fixed at room temperature for 24 hours before being delivered to KAUST

Nanobiology Core Lab facilities for micrography work including cryosubstitution and negative staining.

Storage for future experiments

After several colonies (at least five) were sufficiently isolated per plate, they were picked by the sterile loops described and placed into 0.5 ml dd H2O and 1 ml 30% glycerol solution, in 1.5 ml screw cap tubes for long term storage at -80°C.

Planctomycetes have been known to re-culture after several months storage in glycerol solution, and storage in water is for purposes of future DNA purification and amplification.

64

CHAPTER 3: RESULTS AND DISCUSSION

Results

Within a few days of the processing of the samples, KAUST CMOR successfully identified the seagrass species collected (Fig. 6). The taxonomic diagnosis upon examination of leaf specimens under the microscope were as follows (Batang 2012):

Rabigh Harbor

Halodule uninervis: very distinctive from other species in appearance because of their long, narrow leaves, each with a central vein and trident tip.

Cymodocea serrulata: leaves broader than C. rotundata, with more longitudinal veins

(17 in the specimen at hand); leaf tip is rounded and serrated.

Al Kharrar Lagoon

Halophila stipulacea: paired leaves arising from petioles along the rhizome, with leaves obovate and not narrowing at base; leaf margin spinulose. Along the Saudi Red Sea coast, this species was first recorded by Aleem (1979) at Mugeirma Port (far south of

Jeddah). 65

Figure 9. Seagrass samples collected in Rabigh Harbor (a and b) and Al Kharrar Lagoon (c) with their taxonomic diagnosis displayed. Ruler on left gives length in cm for reference.

Altogether, 51 isolates were purified from C. serrulata seagrass and sediment samples,

52 isolates were purified from H. uninervis seagrass and sediment samples, and 80 isolates were purified from H. stipulacea seagrass and sediment samples. The main findings of this experiment are described in detailed tables with qualitative, subjective notes that were recorded throughout the process of bacterial cultivation. The main objective was to document what the plates looked like in general and if there were any new colonies observed. Once colonies were pure, they were picked and stored immediately thereafter in water and glycerol. The tabulated documentation, though constructed using subjective adjectives, served its purpose of keeping organized the 66 hundreds of plates generated throughout this process. Appendix B contains Tables 2-21 relevant to this section.

After a few days of growth on the agar and gellan media, it became very apparent that bacteria on the plates without antibiotics were much faster-growing than those with antibiotics. Although all plates were checked routinely, it was decided that the 2216 and 1/10 diluted 2216 plates were a priority to screen for isolates over the M13 and anti-2216 plates because of this faster growth rate. Isolates were picked and restreaked on new plates of the corresponding media, with HU standing for H. uninervis, CS standing for C. serrulata, HSN standing for H. stipulacea "normal," and HSA standing for

H. stipulacea abnormal. By the end of April, Tables 2-5 of Appendix B had been constructed, complete with descriptions of all plates inoculated with seagrass biofilm samples, with 1 indicating the sample was undiluted, 2 indicating the inoculum was a

1:5 dilution, and 3 indicating the sample was a 1:25 dilution. Isolates picked from 2216 and 1/10 2216 plates were given updated descriptions in the third column.

Overall, the main trend that can be observed is a decrease in size and number of colonies as the concentration of the inoculum goes down. Oftentimes the morphologies of colonies looked very similar across 1-2216, 2-2216, and 3-2216 of the same inoculum species, but samples preceded by "1-" almost always had greater density of bacterial growth. In general, growth on 2216 plates and 1/10-2216 plates tended to be similar, but the variety of color and shapes of colonies on 2216 plates was overwhelming and completely covered the plates very quickly to the point that within days there was no 67 room for new growth (Fig. 7). The diluted plates gave the slow-growing bacteria a chance to compete and there were sometimes morphologies on 1/10-2216 plates that were not observed on 2216 plates because of the oligotrophic environment they provided.

Figure 10. Photo of 2216 plate inoculated with undiluted H. uninervis biofilm sample. It is representative of the diversity of colony morphology typically seen on 2216 plates after growth of the primary inoculum. Picture taken on 2/5/12.

The plates containing antibiotics had very little growth in the beginning, which was to be expected, but the growth they did have looked very different than the growth on the non-antibiotic plates, with colonies tending to be more convex, glossy, raised, and homogeneous in a pigmented color, either pink, red, or orange. In contrast, by early

May, many of the 2216 and 1/10-2216 plates were threatened by an agar-eating bacterium that acidified the agar substrate as it converted it to a black fluid, with either 68 a different, white phase of the species living on top of the black phase or a different white species living symbiotically with the agar-eating species but displaying aerobic preferences. Ultimately these plates had to be discarded.

Next, the plates spread with sediment inoculum were characterized. Results from all three concentrations (1 = undiluted; 2 = 1:5 dilution; 3 = 1:25 dilution) for sediment collected near C. serrulata are shown in Table 6 of Appendix B. Results from H. uninervis sediment are shown in Table 7, and results from H. stipulacea are shown in Table 8 of the Appendix. The white and black agar-eating organism was much more frequent in the sediments collected from Rabigh Harbor, while H. stipulacea sediment bacteria did have a yellow-colored agar-eating organism but the most commonly seen species was a dull, white, milky bacteria that crawled across the plate. The non-diluted inoculum had too many sediment particles to observe bacteria growth at first because of the consistency of the sample inoculum.

After several days’ growth, many of the enrichment flasks were ready for streaking on the solid media. In nearly every case there were two plates streaked per flask: one from the enrichment broth, and one from the swabbed sludge build-up on the side of the flask. In early May, the plates streaked from flasks were characterized and the results can be found in Tables 9-11 in Appendix B. The growth typically fell into two categories, either white-yellow growth that usually covered the plates in bacterial lawns, or more globular pink-brown, shiny, wet, irregular colonies. Contamination from fungus was also 69 more common in the flask inocula. Appearance of a brilliant orange fungus with white tips was noted for the first time from H. stipulacea-enriched inoculum.

After two weeks growth in enrichment broth, several unusual ball structures (at any given time there were up to seven) were observed in the flasks inoculated with C. serrulata biofilm (undiluted). They ranged from a few mm in length to a few cm, and were almost perfectly spherical. They appeared to be "fuzzy," and when transferred to

M13 agar and gellan plates for growth on solid media, it was observed that these structures were vey resistant to pressure and very resilient, returning to their original form after pressure is removed. Their consistency was almost rubber-like. Their colors ranged from white to gray to light brown. After nine days growth on both agar and gellan plates, however, the three ball structures that were transferred were taken over by fungus; that is, if the structures themselves were not fungal. These results are documented in Table 10 of Appendix B. Growing in conjunction with the balls in the C. serrulata enrichment flask were tiny white rosette-like flakes, and it was hypothesized that these white particles were giving rise to the larger spherical structures. One of these rosettes was also placed on a gellan plate, but was also taken over by fungus.

The ball structures were sent to KAUST Nanobiology Core Lab facilities for electron micrography analyzation, but regrettably they were too large to obtain results. The rosette-forming bacteria, however, were able to be photographed, but poor processing yielded low quality pictures (Fig. 8). 70

Figure 11. Bacteria present in the white rosette-like flakes accompanying the ball-like structures identified in enrichment broth. Sample processed by KAUST Nanobiology Core Lab. Two different organisms of the same sample are represented by a and b; 0.2 µm bar shown in lower left.

Presence of Planctomycetes was suspected on one plate in particular that was inoculated with sludge swabbed from the side of the anti-2216 enrichment broth, which had originally been inoculated with H. stipulacea abnormal biofilm sample. Under the light microscope, motile, teardrop-shaped cells were observed, sometimes in rosette formations, although it is difficult to see in the jpeg files given the constraints of light microscopy (Fig. 9). 71

Figure 12. Specimens from a colony picked on an anti-2216 plate that was inoculated with sludge swabbed from the side of a flask enriching H. stipulacea abnormal biofilm sample. Taken on 5/5/12 with a 40X objective lens. Although not possible to distinguish at this magnification, arrows represent possible rosette formations; a and b are two different frames of the same sample.

The plates streaked using sediment samples enriched by broth (Table 12, Appendix B) also yielded interesting results. After they were re-streaked on corresponding plates, many plates had small, pinpoint, convex colonies with orange or pink pigmentation.

After a preliminary look under an oil immersion 100X objective light microscope, it was suspected that many of these colonies, all shiny orange, pink, or salmon-colored with convex morphology, were Planctomycetes. Samples were prepared in 2.5% glutaraldehyde and sent for TEM, resulting in the figure below (Fig. 13). Additional pictures of the same sample can be viewed in Appendix C. The specific colony picked was grown on an M13 plate, streaked from broth enrichment of sediment collected near

H. stipulacea. 72

Figure 13. Specimen processed from sample grown on an M13 plate which was streaked from broth enrichment of sediment collected near H. stipulacea and observed using TEM after negative staining with uranyl acetate. Complex sheathed flagella and crateriform structures easily discernible; scale bar 200 nm.

Documentation of results continued as plates were checked for several more rounds and isolates collected and either restreaked as additional passages or stored in glycerol and water. Tables 13-16 in Appendix B contain descriptions of the actions taken regarding the isolates from the plates initially inoculated with biofilm. Frequently, isolates that were not yet pure had both a white, shiny, raised, irregular colony with a red or pink pigmented streak threaded through, and these two morphologies had to be re-passaged several times to effectively separate them. After four weeks, the M13 agar 73 and gellan plates along with the anti-2216 plates from the biofilm inocula began to have a significant number of convex, raised colonies with a point (almost "tent"-like) that tended to be shiny and either orange or pink. Although relatively small and pinpoint- sized, there was a sufficient number to re-passage them as well as store them for future use. Thus, by the four week point, the isolates from the first passage of 2216 and 1/10 diluted 2216 plates were virtually all purified or nearly so, and colonies began appearing on the more selective media that had not been observed before, or in great enough number that they could be re-passaged.

The colonies on plates with inocula originating from the enriched flasks began to grow

(Table 17, Appendix B), but very slowly. Often the first passage had to be re-passaged onto less selective media such as 2216 or 1/10-2216 plates because subsequent generations looked less healthy and were growing more slowly. In general, these colonies easily formed microcolonies but had difficulty becoming larger.

When the plates with inocula that had originated from sediment were examined (Table

18, Appendix B), it was discovered that many plates with selective reagents had no to little growth. The majority of the isolates collected from this sub-group came from 2216 and 1/10-2216 plates. They tended to be very moist and mucus-like in consistency, with the smell characteristic of Actinobacteria. Of particular interest was the greater success of the M13 gellan plates compared to M13 agar plates in cultivating colonies. Gellan plates usually grew brown and/or pink colonies that were shiny and convex with a point. 74

Occasionally the M13 agar plates grew orange-brown colonies, but they were fewer in number than the gellan media, and more often than not there was no growth visible.

Finally, for all the remaining isolates that either had been slow-growing or had required additional passages for purification, an exhaustive list was compiled of their descriptions and the action taken in regards to the status of the plate. Table 19 (Appendix B) contains information on isolates collected from plates originating from plates that had been inoculated with biofilm; Table 20 (Appendix B) contains information on isolates collected from plates originating from plates inoculated with enrichment broth or sludge; Table 21 (Appendix B) contains information on plates originating from plates that were inoculated with sediment. By this point, many plates exhibited little to no growth and were kept for observation, but no additional growth was observed even several weeks later.

The 51 isolates purified from C. serrulata samples, 52 isolates purified from H. uninervis samples, and 80 isolates purified from H. stipulacea samples taken together yielded a total of 183 isolates stored in glycerol and 180 corresponding isolates stored in dd H2O at -80°C for future sub-culturing and molecular identification.

Discussion

A number of interesting results arose over the course of the cultivation process. From start to finish, not only were the tabulated notes recorded to document the progress of 75 the isolates, but predicted and unexpected observations were made along the way. The predicted result was that the isolation methods employed would indeed capture

Planctomycetes. The unexpected developments were the spontaneous growth of the ball-like structures and the associated white flakes suspended in solution.

After inspecting a number of colonies displaying morphological features characteristic of

Planctomycetes using light microscopy, we had reason enough to examine the specimens in more detail using electron microscopy. The results obtained from TEM further strengthened our belief that the colonies in question do indeed represent

Planctomycetes (Fig. 10 and Appendix C) and further molecular work will confirm or disprove these suspicions.

Easily discernible in Figure 13 are the crateriform structures so distinctive of

Planctomycete species. The polar distribution of crateriform structures marks this as a

Pirellula-like strain, a suspicion confirmed through personal communication by Dr. Lafi with Dr. Fuerst, a world-renowned Planctomycetes expert. The complex sheathed flagella is another trait commonly found in Pirellula strains versus other Planctomycete species. Other negatively stained Pirellula cells in Fig. 11b and 11c can be compared to the cells processed from H. stipulacea biofilm, and additional pictures of the sample can be viewed in Appendix C. The structural features of the unidentified strain and the

Pirellula strains from previously published work are strikingly similar. 76

Figure 14. Comparison of suspected Pirellula species with confirmed Pirellula strains a) Same specimen as that of Figure 13 from Results; included for comparison’s sake with b) IFAM 1313 strain later re-named Pirellula marina from Schlesner (1986) and c) strains of Pirellula from Fuerst (1997) as follows: 1. non-pigmented AGA/M12 strain 2. pink- pigmented HGG/MPI strain 3. orange-pigmented AGA/M18 strain 4. Planctomyces brasiliensis. Scale bars for a and b as shown; thin lines in c 1-4 indicate 0.5 μm. Flagella are labeled (fl) as are crateriform structures (cr).

77

The unexpected presence of the ball-like structures is still a mystery. Even after the conclusion of this study, they continued to arise spontaneously from seemingly clear broth, always accompanied by the white flakes which TEM had shown are associated with a bacterial species (Fig. 8). Without molecular identification it cannot be confirmed that these were Planctomycetes, but it is certainly possible, given that their life-cycle is divided between juvenile motile and mature sessile phases. If these had indeed been

Planctomycetes cells, we could have been observing juvenile cells free in aqueous solution under the microscope, and it is possible that as they matured from free-living to aggregate-forming they developed into colonies inhabiting or composing the enigmatic ball structures, uninhibited by other non-beta-lactam-resistant bacteria and free to flourish in the flask with plenty of Planctomycetes-specific nutrients.

As to why this sort of macrostructure would not have been observed on the solid media, it may be related to the reason that the inocula from enriched broth never gave rise to any growth larger than microcolonies. In fact, it can even be expected that bacteria in inocula originating from suspended enrichment in an aqueous environment would have a difficult time adapting to reproduction on solid media substrate. This could explain why the first passage was so much more successful than subsequent passages. Clone libraries have had more success documenting Planctomycetes that live as marine aggregate attached bacteria than they have free-living bacteria (Bengtsson and Ovreas

2010). Therefore, problems arising from using different phases of media had been anticipated, and were in fact encountered. 78

Otherwise, very few unforeseen issues arose during the course of the experiment.

Occasionally, the size or number of colonies was too small for storage in both glycerol and water. In these cases, because priority was given to sub-culturing for future experiments, storage in glycerol only was chosen. After these isolates are re-grown, 16S rRNA amplification can be performed to ascertain identity.

The plan was to document all colony formation and morphology, take note of bacteria suspected to be Planctomycetes, and investigate them further to confirm their identity.

The other bacteria isolated are of interest from an ecological perspective, but they are certainly not representative of all the bacteria from the original inocula, and isolation techniques are not meant to measure biodiversity. Although characterizing the microbial community is beyond the scope of this study, it is a logical next step in understanding Planctomycetes and the role they play in microbial communities.

Overall, 183 different isolates were cultivated and purified using media that we hypothesized would be able to isolate Planctomycetes, and light and electron microscopy were able to confirm this. Future experiments will need to involve 16S rRNA gene amplification to definitively identify the isolates, but for the purpose of this study, morphological and TEM analysis were sufficient.

79

CHAPTER 4: CONCLUSION

The goal of this study was to meet the research objective and hypotheses outlined in the Introduction section. The first hypothesis was that Planctomycetes could be found in the biofilm of Red Sea seagrasses. In order to test this hypothesis, confirmation of the second hypothesis was first necessary because confirmation of their presence first required that conventional enrichment and isolation methods be employed for cultivating the bacteria.

Enrichment broth and solid media were prepared in advance of the collection of seagrass and sediment samples. There were three types of broth - M13, anti-2216, and

1/10-2216 - and five types of plates - M13 agar, M13 gellan, anti-2216 agar, 1/10 2216 agar, and 2216 agar - used in the attempt to isolate Planctomycetes. The media containing selective antibiotic and antifungal agents successfully prevented domination by fast-growing bacterial species, and allowed time for slow-growing bacteria to colonize. After several weeks, each plate type had colonies with a distinct set of morphological features. By the end of the study, the only suspected Planctomycetes identified were grown on gellan and agar M13 media, with gellan being the slightly more successful media, although it is possible that some Planctomycetes grew on anti-

2216 plates. Polymerase chain reaction (PCR) gene amplification will identify which isolates are in fact Planctomycetes strains.

The third hypothesis stated that pure culture of Planctomycetes, confirmed through morphological study and electron microscopy, could be obtained, and the cells used for 80 further experimentation. TEM work did indeed produce satisfactory results, in line with the caliber of results produced in the field of Planctomycetes microbiology (Schlesner

1986; Fuerst, Gwilliam et al. 1997). The morphology of the isolated and purified cells is consistent with previous findings (Fuerst 1995; Lindsay, Webb et al. 2001), and their proper storage in glycerol has ensured that they may be sub-cultured for future planned experiments. Because of the confirmation of the third hypothesis, we know that the second was also confirmed because the isolation techniques utilized proved appropriate for the study of Red Sea seagrasses. The confirmation of the second hypothesis, in turn, supports the first hypothesis, because the cultivation methodology was applied to examination of Red Sea seagrass biofilm. The results demonstrate the viability of all three hypotheses, and so we can state that the objective of this study has been met.

The discovery of the presence of Pirellula species in the Red Sea is significant if only for the reason that Planctomycetes have never been cultivated from samples collected from the Red Sea before. The Saudi Red Sea coast has been studied almost strictly from a sedimentological perspective up until the last three years (El-Sayed 1987; Washmi

1999; Basaham 2008). Knowledge pertaining to the specific micro-organisms that are present and how they are contributing to ecological cycles is nearly non-existent (Viney

2011). This finding adds to the scientific body of work on Planctomycetes, as well as Red

Sea microbiology and ecology. By achieving a better understanding of how

Planctomycetes interact with the seagrass biofilm from which they were cultivated, contribution is made not only to Red Sea marine science, but scientific knowledge 81 regarding prokaryotic and eukaryotic relationships in general (Bengtsson and Ovreas

2010). By overlooking the system of this ecological niche, gaps have been permitted in the collective understanding of how Planctomycetes contribute to the global carbon, sulfur, and nitrogen cycles. Part of closing these gaps is searching for where

Planctomycetes are ecologically key players. Further experimental work must be carried out to investigate the interactions of Planctomycetes with their environment. Genomic analyses of Planctomycetes have laid the groundwork in identifying which biochemical pathways to focus on (Jenkins, Kedar et al. 2002; Bauer, Lombardot et al. 2004), but their metabolism and the significance of their compartmentalized structures are still vastly understudied, with perhaps the exception of anammox (Jetten, Sliekers et al.

2003).

Successful cultivation of Planctomycetes is only the initial phase. Next comes the molecular work in identifying the isolated bacteria through PCR amplification. Genomes need to be added to the limited Planctomycetes databases, a feat achievable through pyrosequencing. FISH can be utilized in conjunction with Planctomycetes-specific probes (Pizzetti, Fuchs et al. 2011) to examine environmental samples and the positioning of Planctomycetes in those samples, and terminal restriction fragment length polymorphism (TRFLP) can profile the microbial community of the niches where they are found (Bernhard, Marshall et al. 2012). In the immediate future, the next step is to sub-culture the Planctomycetes that are currently stored in glycerol at -80°C, and run various enzyme assays to test for activity. Transcriptomic work can be carried out 82 simultaneously to elucidate different pathways by characterizing the functionality and expression levels of the genes encoding for the enzymes involved. This would add to the collection of experiments already dedicated to this topic (Pearson, Budin et al. 2004;

Wecker, Klockow et al. 2009), generating data that could be used for industrial and ecological application.

Cataloging and amassing data on hundreds of isolates is an essential first step in the foundation of future studies that can accelerate the pace at which Planctomycetes revolutionizes industry and scientific understanding of global nutrient cycling.

83

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APPENDICES

Appendix A: TEM Protocol

The following protocol was used by Dr. Rachid Sougrat from KAUST Nanobiology Core

Lab to process samples (Sougrat 2012).

Negative staining on whole individual cells:

Apply 2 to 4 µl of solution containing bacterial cells to the grid for 30 s and blot with filter paper.

Rinse the grid with a few drops of uranyl acetate (1%) and blot the excess with filter paper.

Sections on pellets/aggregates:

Cells were fixed with 2.5% glutaraldehyde in 0.1 M cacodylate buffer, pH 7.4. Cells were washed and post-fixed for 1 h at room temperature in reduced osmium (1:1 mixture of

2% aqueous osmium tetroxide and 3% aqueous potassium ferrocyanide). After fixation, cells were pre-embedded in agar (2%), dehydrated in ethanol, and processed for Epon embedding. Thin (80-nm) sections were cut and collected on a copper grid and contrasted for 2 minutes with lead citrate. Sections then were examined with a Titan

80-300 electron microscope operating at 200 kV and equipped with a 4k × 4k charge- coupled device (CCD) camera (Gatan, Inc).

Appendix B: Tabulated qualitative descriptions of colony morphology

Tables 2-21 contain all results documented regarding colony morphology are grouped by category according to their titles for easy access to information. A number of 93 different abbreviations are used, and in some cases included in the table description for clarity, but the following list should be sufficient for understanding the shorthand used at the time of recording:

M13: agar M13 plate

GM13: gellan M13 plate anti-2216: 2216 plate plus antibiotics

1/10-2216: 2216 plate diluted 1/10

CS: C. serrulata

HU: H. uninervis

HS: H. stipulacea

HSN: H. stipulacea “normal”

HSA: H. stipulacea abnormal

1: original inoculum was undiluted

2: original inoculum was diluted 1:5

3: original inoculum was diluted 1:25 sed: sediment

B: broth

S: sludge

F: flask

94

Table 2. Description of colonies growing from all three concentrations of H. uninervis inoculum on all five media types as of 29/4/12, with updates on isolates that were picked previously Sample inoculum concentration Progress of isolates and plate type Qualitative description of plate picked earlier, if any ½ plate speckled with black mold/fungus; ½ covered with brown mold/fungus (brown = 1-GM13 ring-like); 1 white colony present Green-brown fungus and 1-2 1-M13 white, 2 orange, 1 black bacterial colonies present Black fungus with a few pink 1-anti-2216 bacterial colonies HU7 = deep red streak; HU8 = mostly white 1-1/10-2216 Many white and pink colonies streaks with yellow and rose pigmentation; not pure HU1 = white and orange streaks; HU2 = orange, shiny, circular; HU3 = Very multi-colored colonies – black; HU4 = deep red black, pink, yellow, white, green, and beige swirls; HU5 = orange, etc. See Figure 9 deep red, circular, large, glossy, plus light pink with same 1-2216 characteristics 2-GM13 mostly white, some pink, 1 black 2-M13 1 pink many small pink, medium white, 1 very large orangeish, slightly 2-anti-2216 raised HU13 = rose and white streaks; HU14 = rose multi-colored, small colonies - circles, smaller; HU15 = pink, yellow, purple, orange purple, mostly pure; HU16 = white, large, 2-1/10-2216 irregular 95

HU9 = some deep pink, irregular, surrounded by white, others lighter purple, circular, still surrounded by white; HU10 = range from small multi-colored, small - black, pink, circles to irregular yellow, orange, purple blocks, raised orange but not one color - layers; HU11 = white w/ pink hue, glossy, slightly raised; HU12 = some deep purple, rest 2-2216 irregular white 3 orange-tinged/beige colonies, 3-GM13 glossy, raised 2 small beige, 1 small pink, raised, 3-M13 glossy 3-anti-2216 2 small pink, convex HU17 = very small beige; HU18 = very small deep multi-colored, mostly yellow, red, raised; HU19 = orange, pink coffee-colored-yellow small, irregular; HU20 - 3-1/10-2216 light pink/beige swirls HU21 = deep pink, very concentrated, raised, glossy; HU22 = whitish- large, multi-colored; mostly beige + purple, pink, yellow yellow, large; HU23 = yellow-orange, beige, and deep pink; HU24 = 3-2216 small light yellow-dotted

Table 3. Description of colonies growing from all three concentrations of C. serrulata inoculum on all five media types as of 29/4/12, with updates on isolates that were picked previously Sample inoculum concentration and plate Progress of isolates picked type Qualitative description of plate earlier, if any 96

1-M13 1 pink-beige, large; 1 very small 1-GM13 fungus-infested lots of fungus, a few white 1-anti-2216 bacterial colonies CS3 = white and pink, small; CS4 = similar to CS3 but more pink; small, multi-colored, mostly white, CS9 = tiny, yellow-orange; CS10 1-1/10-2216 some pink, orange, yellow = small white CS7 = large, irregular, light pink, raised, glossy; CS8 = similar to CS7, slightly smaller colonies; CS1 = large, irregular, light pink; CS2 = same as CS1, slightly more orange; CS5 = pink and white, large, irregular; CS6 = black, many small white w/ some small raised, spread through bottom 1-2216 yellow/black of agar 2-M13 2 medium-sized light orange fungus, 1 pink bacterial colony at 2-GM13 edge 2-anti-2216 pink, tiny, circular; 1 large beige CS15 = white and orange, scattered white/beige; 1 pink- irregular; CS16 = light pink, very 2-1/10-2216 orange similar to CS15 CS11 = amber, raised, irregular, glossy; CS12 = 1 vibrant pink, the rest large and light pink; CS13 = multi-colored, black, purple, beige, white, circular; CS14 = same as 2-2216 yellow CS13 3-M13 no growth visible 3-GM13 no growth visible 3 places with pink colonies raised, 3-anti-2216 glossy CS17 = mostly tiny white with some pink; CS18 = vibrant pink, clustered, irregular distribution of pigmentation; CS23 = orange multi-colored yellow, orange, and white blended, glossy, 3-1/10-2216 white raised; CS24 = white 97

CS19 = vibrant pink, large, irregular, and light pink/white; CS20 = white, small, some with deep black and watery center; CS21 = pretty orange, very mostly white, medium-large, and homogeneous color; CS22 = 3-2216 black, pink, musk, yellow pretty yellow, very homogenous

Table 4. Description of colonies growing from all three concentrations of “normal” H. stipulacea inoculum on all five media types as of 29/4/12, with updates on isolates that were picked previously Sample inoculum concentration and plate Progress of isolates picked earlier, type Qualitative description of plate if any 3 small circular white colonies, 1-M13 convex strikingly similar to M13 but 3x 1-GM13 the colonies 1-anti-2216 no growth observed HSN1 = transparent - looks like water droplets/dew; some have pink dot in center; HSN2 = orange- yellow, small streaks; HSN3 = similar to HSN2 but w/ 2 distinct, separate colonies; HSN4 = similar small transparent white and to HSN1- fast-growing, over 1/2 of 1-1/10-2216 yellow-speckled entire plate HSN5 = irregular, glossy, raised white, one pink area; HSN6 = pink-orange colonies surrounded by lighter white area, some areas have "pointy" corners; HSN7 = pink/rose white, raised, glossy; HSN8 = same as HSN7 but w/ covered in white and yellow pink; HSN9 = large, globbed colonies, some circular, some together white colonies, some w/ irregular, a few pink/deep flecks of brown; HSN10 = white, 1-2216 purple glossy, thin lines, some tinged red 98

1 large orangeish circle-ish flat but raised at center, some 2-M13 others similar but smaller 2-GM13 13 small white convex 1 pink small glossy; fungus 2-anti-2216 infested - dime-size clear/white small growth covering plate; yellow patches of clustered colonies that appear to HSN21 = flat yellow glossy be eating into agar, forming patches; HSN22 = same as HSN21 2-1/10-2216 divots but stronger orange color covered in whiteish, clear lawn HSN19 = happily growing orange w/ dots of yellow; resembles streaks; HSN20 = white, small, 2-2216 "veins" circular 3-M13 3 small yellow-white convex fungus-infested; 1 small beige 3-GM13 colony on side 3-anti-2216 no growth visible mostly irregular white/clear colonies of various sizes all over plate; one side has yellowish HSN11 = no growth visible; HSN12 3-1/10-2216 colonies eating into it = no growth visible HSN13 = fat globby orange-pink- white homogenous; HSN14 = rose-white swirls growing in concentric circles; HSN15 = intense red-orange surrounded by thin transparent layer, flat; HSN16 = white, fat, glossy, raised; HSN17 = 1/2 yellow, 1/2 white-rose, more/less circular alternating 2x; HSN18 = raised white/transparent colonies, black, black inhabiting agar all the 3-2216 some rose, yellow, deep black way through

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Table 5. Description of colonies growing from all three concentrations of abnormal H. stipulacea inoculum on all five media types as of 29/4/12, with updates on isolates that were picked previously Sample inoculum concentration and plate Progress of isolates picked earlier, type Qualitative description of plate if any 1-M13 1 very small white colony 13 very small orange white 1-GM13 convex 1-anti-2216 no growth visible white-yellow lawn, very HSA18 = mostly white, irregular, textured, areas where yellow are speckled - some w/ rose; HSA19 = 1-1/10-2216 biting into agar same as HSA18 HSA20 = clumpy irregular white, raised, pretty red, specks throughout; HSA21 = orange- yellow streaks, fade into white at one end; HSA22 = black agar- eating colonies, white milky surfaces with black sinking to bottom, so far only halfway down; HSA23 = flat streaky tossed b/c taken over by black orange-red, in some places dull, colonies eating through agar; interspersed/alongside small 1-2216 byproduct = aqueous, acidic fluid circular brown 2-M13 no growth visible 2-GM13 14 orange-yellow convex 2-anti-2216 1 small yellow colony visible HSA1 = transparent, white streaks, single colonies, basically white transparent, some yellow, circular; HSA2 = clear globby fat causing more texture; 1-2 irregular; HSA3 = white, flat w/ 2-1/10-2216 orangeish transparent irregular deep red in center - small points 100

HSA4 = orange, glossy, surrounded by white areas, raised/pointy texture in deep orange parts; HSA5 = same as HSA4; HSA6 = deep red circular, raised, bright red middle surrounded by white; HSA7 = white irregular globby pinkish; HSA8 = mostly white, transparent, chunky, glossy, some pink-red splotches; HSA9 = small orangeish film coating plate; one circles, white-rose homogeneous; section being eaten away; one HSA10 and HSA11 = virtually 2-2216 whitish path, a few red circles identical to HSA8 3-M13 no growth visible 3-GM13 2 orange-white colonies visible weird yellow "venous" growth in 3-anti-2216 center; highly irregular HSA16 = pink small points surrounded by white, some pure hard-to-see white transparent white; HSA17 = irregular raised 3-1/10-2216 growth covering plate white HSA12 = no growth visible; HSA13 = 2 irregular light orange globs, raised, shiny; HSA14 = some glossy globs, fat, rose-white white globby growth in lawn; homogenous, interspersed w/ some musky rose colonies; one smaller intense yellow, shiny 3-2216 black irregular patch circles

Table 6. Description of colonies growing from all three concentrations of C. serrulata sediment inoculum on all five media types as of 1/5/12; no isolates picked at this point Sample inoculum concentration and plate type Qualitative description of plate no growth visible; fungus infested; possibly one small white colony 1-M13 but hard to tell if artifact 1-GM13 1 white-orange, small colony 101

growth only visible in areas pockmarked w/ tiny bubbles - one 1-anti-2216 section = orange, one section = pink covered in transparent white irregular growth; shiny; raised just 1-1/10-2216 slightly black/white agar-eating organism has completely taken over plate - texture of surface completely uneven and some areas have small projections of white growth while others have black all the way to 1-2216 bottom of plate 2-M13 no growth visible 2-GM13 no growth visible 2-anti-2216 no growth visible milky white transparent white colonies covering plate; one yellow 2-1/10-2216 patch slightly raised; dew-like reflection 2-2216 fungus-infested; tossed 3-M13 no growth visible 3-GM13 no growth visible 3-anti-2216 no growth visible milky white transparent globby lawn; some circular colonies, some 3-1/10-2216 fat globs black and white agar eating; wider circles than 1-2216 or 2-2216; 3-2216 higher ratio of white to black

Table 7. Description of colonies growing from all three concentrations of H. uninervis sediment inoculum on all five media types as of 1/5/12; no isolates picked at this point Sample inoculum concentration and plate type Qualitative description of plate 1-M13 fungus starting to grow; no growth visible 1-GM13 1 orange circular convex; another of same starting nearby 1-anti-2216 no growth visible 1-1/10-2216 small white speckled colonies dotted all over large orange flat irregular colonies varying in size; some areas white, 1-2216 some black 2-M13 no growth visible 2-GM13 fungus infested; no growth visible 2-anti-2216 no growth visible milky white, translucent, irregular; one irregular shiny patch w/ 2-1/10-2216 orange in center; some patches have yellowish hues 102

one patch goes down center, very dull, rough texture; on either side 2-2216 are pockets digging into agar 3-M13 no growth visible 3-GM13 fungus starting; no growth visible 3-anti-2216 no growth visible 3-1/10-2216 milky translucent white, irregular, sort of reflective orange eating into agar; few black dots on perimeter, white clearish 3-2216 in remaining area, all shiny

Table 8. Description of colonies growing from all three concentrations of H. stipulacea sediment inoculum on all five media types as of 1/5/12; no isolates picked at this point Sample inoculum concentration and plate type Qualitative description of plate 1-M13 hard to distinguish sediment particles from bacterial growth 1-GM13 hard to distinguish sediment particles from bacterial growth 1-anti-2216 hard to distinguish sediment particles from bacterial growth 1-1/10-2216 milky translucent white; irregular; sort of reflective orange-eating into agar; few black dots on perimeter, white clearish 1-2216 in remaining area; all shiny 2-M13 no growth visible 2-GM13 no growth visible 2-anti-2216 no growth visible 2-1/10-2216 2 deep purple agar-eating; rest = transparent/white splotches 2-2216 white, milky, translucent all over; little to no variation in color 3-M13 no growth visible 3-GM13 no growth visible 3-anti-2216 dull, white, almost not visible colonies, but unclear if bacteria 3-1/10-2216 milky white transparent, irregular, shiny globs 3-2216 yellow agar-eating; textured surfaces; yellow/white/rosy patches

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Table 9. First description from 1/5/12 of plates streaked using inocula from H. uninervis biofilm samples that were first enriched in selective media; B = the plate was streaked with broth directly; S = the plate was streaked with sludge swabbed from the side of the flask Sample inoculum and plate type Qualitative description of plate FHU-M13-B fungal growth - spicules, raised, spiky, in isolated dots covering plate FHU-M13-S fungal growth - spicules, raised, spiky, in isolated dots covering plate FHU-GM13-B fungal growth - spicules, raised, spiky, in isolated dots covering plate FHU-GM13-S fungal growth - spicules, raised, spiky, in isolated dots covering plate FHU-anti- 2216-B dull, textured white lawn FHU-anti- 2216-S thick lawn, white-yellow; no distinct colonies FHU-1/10- dew-like, glistening, irregular growth, separated, large "droplets" 2216-B interspersed throughout smaller more irregular growth FHU-1/10- dew-like glistening irregular growth patterned in small lines; 2216-S transparent FHU-1/10- white milky blob covering almost entire center - 50% area - rest = 2216-B spotty texture; dull FHU-1/10- 2216-S same as broth but w/o white patches; till there but in streak form

Table 10. First description from 1/5/12 of plates streaked using inocula from C. serrulata biofilm samples that were first enriched in selective media; B = the plate was streaked with broth directly; S = the plate was streaked with sludge swabbed from the side of the flask Sample inoculum and plate type Qualitative description of plate FCS-M13-B fungus-infested; no bacteria FCS-M13-S fungus-infested; no bacteria FCS-GM13-B fungus-infested; no bacteria FCS-GM13-S fungus-infested; no bacteria FCS-anti- 2216-B dull, textured white lawn FCS-anti- 2216-S pinkish brown, very raised growth; shiny, irregular FCS-1/10- 2216-B white, transparent, irregular, shiny, growth with some white colonies 104

FCS-1/10- 2216-S white lawn, shiny and dull brownish lawn covering everywhere except a previously scraped FCS-2216-B rectangle; some areas have brown circles; hard to tell color FCS-2216-S same as broth but with more, smaller circles; brown seems darker FCS-GM13- rosette fungus; no bacteria FCS-M13- ball1 fungus; no bacteria FCS-M13- ball2 fungus; no bacteria FCS-GM13- ball3 fungus; no bacteria

Table 11. First description from 1/5/12 of plates streaked using inocula from H. stipulacea biofilm samples that were first enriched in selective media; B = the plate was streaked with broth directly; S = the plate was streaked with sludge swabbed from the side of the flask Sample inoculum and plate type Qualitative description of plate FHSN-anti- 2216-B small pink colonies, spread evenly and distributed everywhere FHSN-anti- very tiny small pink dots coating surface in haze; one area = orange; 2216-S one black irregular raised lump mostly large pink/brown colonies dotting plate, irregular, raised but FHSN-1/10- flat, "serrated" undulating edges; also smaller section w/ white 2216-B circular colonies FHSN-1/10- brown small colonies, all oriented (ovoid) in direction of streak - fuso- 2216-S form - maybe indicative of low oxygen tolerance FHSA-anti- overtaken by brilliant orange fungus - orange has spread entirely 2216-B through agar, white on top; not bacterial growth seen FHSA-anti- dotted with pink, circular, convex/spheres; one corner has orange 2216-S slimy irregular growth; a few black spheres of fungus FHSA-anti- one small pink colony inhabiting bubble; ball seems to be coating w/ 2216-ball fungus feathery, dull texture, white/transparent growth across whole plate, FHSA-1/10- interspersed with tiny spots of no growth zones surrounding dull 2216-B yellow colonies FHSA-1/10- 2216-S orange-yellow dull lawn; one small spot of fungus 105

Table 12. First description from 1/5/12 of plates streaked using inocula from sediment samples associated with all three seagrasses that were first enriched in selective media; B = the plate was streaked with broth directly; S = the plate was streaked with sludge swabbed from the side of the flask Sample inoculum and plate type Qualitative description of plate FHS-sed- many tiny orange-salmon dots, ball-like/convex, shiny; some same M13-B but white FHS-sed- GM13-B many tiny orange-salmon dots, ball-like/convex, shiny FHS-sed-anti- many tiny orange-salmon dots, ball-like/convex, shiny; some same 2216-B but white FHS-sed-anti- 2216-S one single white colony, convex, shiny white, dull, textured film coating surface; many yellow colonies - FHS-sed- pinpoint-sized - scattered throughout, w/ no growth zone 1/10-2216-B surrounding them FHS-sed- orange-brown, dull growth in streaks across plate; several irregular 1/10-2216-S white present as well, flat and dull FCS-sed-anti- 2216-B small, light pink, flat, dull FCS-sed-anti- 2216-S small orange and pink dots, burrowed into pre-existing holes FCS-sed- surface dull, textured; scattered w/ dew-looking colonies; irregular, 1/10-2216-B small, raised FCS-sed- hard to distinguish possible sediment residue from bacterial growth; 1/10-2216-S spots reflective/shiny FHU-sed- 2 white/yellow textured growing in concentric circles; 3 rose-ish anti-2216-B circles, patchy; some pink and white small colonies on outskirts FHU-sed- anti-2216-S orange circles, irregular; white, dull, textured growth FHU-sed- white, shiny, translucent film; some big patches, some spotty 1/10-2216-B colonies; irregular FHU-sed- brownish soil-looking build-up along trails left by inoculation loop; 1/10-2216-S hard to tell if residue or growth

106

Table 13. Second look on 7/5/12 at C. serrulata plates. Inoculum from 1 = undiluted biofilm sample, 2 = 1:5 dilution, and 3 = 1:25 dilution; corresponding re-passaged isolates are described and either re-passaged or stored in glycerol (G) or water. Sample inoculum concentration and plate Description of isolates picked type Qualitative description of plate earlier and action taken 1-M13 fungus 1/2 colony picked and dispersed on other 1/2 of plate; remaining 1-GM13 1/2 spread on anti-2216 1-anti-2216 fungus CS3 = some colonies have flat, shiny pink streaks, some pure white, restreaking as CS3a and CS3b; CS4 = pink streak lines, very pigmented, surrounded by clusters of translucent "dew drops", restreaking as CS4a and CS4b, pink and translucent; CS9 = small yellow-orange circular flat, iridescent - stored G/water; CS10 = white, 'raised', shiny irregular - 1-1/10-2216 stored G/water CS1 = pink-orange irregular growth, shiny, irregular, raised - stored G/water; CS2 = same as CS1 - stored G/water; CS5 = chunky/globby shiny irregular, some pink some white, restreaking as CS5a and CS5b; CS6 = black agar-eating, some lumpy, raised, dull, some shiny - stored G/water; CS7 = light pink-brown, shiny, raised, homogeneous - stored G/water; some of CS7 had blackish thread running through it, restreaking as CS7b; CS8 = white/orange, raised, shiny, 1-2216 irregular - stored G/water 107

2 pink orange circular colonies, raised w/ a point, shiny; spreading 1/2 around remainder of plate, 1/2 on 2216; storing 2-M13 one in G as "CS2 susp. Pla" mostly fungus; one white- orange shiny convex surrounded by fungus; picking 1/2 and restreaking on new GM13 plate, 2-GM13 other 1/2 on 2216 2 kinds of colonies, one pink- coral, round, convex, shiny, small; one flat/raised, shiny, restreaking as CS25 and CS26, 2-anti-2216 orange, larger respectively CS11 = yellow-brown, iridescent, raised, irregular, shiny - stored G/water; CS12 = white and pink raised, shiny, irregular, restreaking as CS12a and CS12b; CS13 = white, raised, flat, shiny - stored G/water; CS14 = orangeish pink lumpy shiny raised irregular - stored G/water; CS15 = orange, irregular, shiny but textured - stored G/water; CS16 = pink-coral, shiny, raised, irregular - 2-2216 stored G/water 3-M13 no growth visible 3-GM13 no growth visible 4 pink-coral raised convex shiny; restreaking as CS27 and CS28, 3-anti-2216 1 orange raised convex shiny respectively CS17 = very small pink dots, raised shiny irregular, restreaking P3; CS18 = pink and white streaks, 3-1/10-2216 restreaking as CS18a and CS18b 108

CS19 = deep red, fly but raised, irregular, large - stored G/water; CS22 = intense yellow, raised, textured, streaky - stored G/water; CS20 = white raised shiny irregular - stored G/water; CS21 = burnt orange/glowing, shiny raised irregular - stored G/water; CS23 = orange shiny raised textured - stored G/water; CS24 = white and yellow mixed, shiny raised irregular, restreaking 3-2216 as CS24a and CS24b

Table 14. Second look on 7/5/12 through 9/5/12 at H. uninervis plates. Inoculum from 1 = undiluted biofilm sample, 2 = 1:5 dilution, and 3 = 1:25 dilution; corresponding re- passaged isolates are described and either re-passaged or stored in glycerol (G) or water. Sample inoculum concentration and plate Qualitative description of Description of isolates picked type plate earlier and action taken fungus-infested; 2 small orange convex round; 1 black round pinpoint shiny convex; 1 white round creating P2 isolate HU25, HU26, 1-M13 pinpoint shiny convex and HU27, respectively 1-GM13 1 round sphere white shiny restreaking as P3 HU28 some buried in bubbles, pink-coral; others in fungus; 1 orange-white convex pink restreaked as HU29; orange 1-anti-2216 shiny; restreaked as HU30 HU8 = 1/2 white, 1/2 yellow, shiny raised irregular, restreaking as HU8a and HU8b P3; HU7 = red and white growing together, irregular shiny, raised, restreaking as P3 1-1/10-2216 HU7a and HU7b 109

HU1 = 1/2 white, 1/2 yellow raised shiny irregular, restreaking as HU1a and HU1b; HU2 = raised textured orange-yellow irregular - stored G/water; HU3 = no growth visible; HU4 = deep red and white, very distinct, storing as HU4a - red, 1-2216 and HU4b - white in G/water 5 circular white convex w/ one point forming 'tent'; 1 storing as HU39 - G/water; same morphology except restreaking as HU38 on 2216 and 2-M13 pink M13, respectively white shiny raised some circular, some crenulated; restreaking as HU35, HU36, and one irregular black; one HU37, respectively, on 2216 and 2-GM13 orange circular shiny GM13 one crenulated textured/shiny, large growth; several orange restreaking on anti-2216 and 2216 shiny raised; smaller pink- as HU31, 32, 33, and 34, 2-anti-2216 coral convex circular respectively HU13 and HU14 = pink circular flat thin dull w/ white growing in and around, restreaking on 2216 as HU13 and HU14; HU15 = purple flat shiny circular - stored G/water; HU16 = flat raised white/transparent, irregular - 2-1/10-2216 stored G/water HU9 = shiny raised brown surrounded by white and pink, surrounded by white - restreaking as HU9a, 9b; HU10 = small dull raised yellow-orange - stored G/water; HU11 = some orangeish, even, "fade into" shiny white irregular raised - restreaking as 12a 2-2216 and b restreaking orange on new M13 3 orange-white circular plate as HU41, one on 2216; shiny convex; one same restreaking pink - 1/2 on M13, 1/2 3-M13 except coral on 2216 as HU42 110

storing some - G/water - restreaking rest around same plate 3-GM13 3, same as HU41 as HU40 several small pink convex restreaking pink on rest of plate coral shiny; one same and on 2216 as HU43; black on 3-anti-2216 except black 2216 as HU44 HU17 = light brown speckled shiny raised circular - stored G/water; HU18 same except deep pink - stored G/water; HU19 = yellow and white, shiny raised irregular, restreaked as HU19a and b; HU20 = pink raised shiny irregular - 3-1/10-2216 stored G/water HU21 = deep pink, thick, raised circular dull - stored G/water; HU22 = white yellow shiny textured raised, restreaking as HU22a and b; HU23 = irregular, shiny, raised, restreaking pink as HU23a, white as HU23b, and yellow as HU23c; HU24 = small pinpoint colonies, all yellow shiny 3-2216 raised - stored G/water

Table 15. Second look on 9/5/12 and 10/5/12 at H. stipulacea abnormal plates. Inoculum from 1 = undiluted biofilm sample, 2 = 1:5 dilution, and 3 = 1:25 dilution; corresponding re-passaged isolates are described and either re-passaged or stored in glycerol (G) or water. Sample inoculum concentration Qualitative description of Description of isolates picked earlier and plate type plate and action taken 1-M13 no growth visible many white-orange convex stored in G/water as HSA24; 1-GM13 raised w/ point shiny restreaking some on GM13 and 2216 1-anti-2216 111

HSA18 = white raised shiny irregular, points and specks of brown, restreaking as HSA18a and b; HSA19 = beige raised shiny irregular - stored 1-1/10-2216 G/water HSA20 = a- round shiny raised circular, b-white-yellow dull textured flat, c- rose flecks within b - restreaking on 2216; HSA21 = yellow orange raised shiny irregular - stored G/water; HSA22 = mucus agar-eating brown and black, restreaking as HSA21 a and b; HSA23 = a-yellow-orange, raised shiny irregular, b-brown shiny raised 1-2216 irregular - restreaking P3 2-M13 no growth visible many orangeish circular stored G/water as HSA25; also 2-GM13 convex shiny, w/ point restreaking on 2216 and GM13 crenulated transparent flat dull; one very small pink 2-anti-2216 convex shiny restreaking on 2216 as HSA26 and 27 HSA1 = white transparent circular raised - stored G/water; HSA2 = flay shiny irregular white - stored G/water; HSA3 = flat shiny irregular red and white, restreaking on 2216 as HSA3a 2-1/10-2216 and b HSA4 and 5 = yellow orange small dot raised shiny irregular, surrounded by clear-white - restreaking as HSA4 - yellow and HSA5 - clear; HSA6 = deep red round/irregular shiny raised/flat - stored G/water; HSA7 = white-pink globby shiny irregular raised - stored G/water; HSA8a = same as HSA7, b is pink swirl inside a, restreaking; HSA9 = same as HSA7 - stored G/water; HSA10 = flat white shiny irregular; HSA11 = orange-white shiny raised mixed w/ 2-2216 brown and pink, re-passaging 3-M13 no growth visible 112

2 orange-pink convex w/ point shiny raised; restreaking one on same 3-GM13 plate, one on 2216 named HSA28 crenulated irregular raised white/yellow, dull and 3-anti-2216 textured restreaking as HSA29 HSA16 = raised shiny circular irregular, mixed pink dots w/ white blobs, restreaking as HSA16a and b; HSA17 = white irregular shiny raised - stored 3-1/10-2216 G/water HSA12 = no growth visible; HSA13 = 2 brownish raised irregular shiny - stored G/water; HSA14 = raised shiny irregular white/yellow mixed, restreaking onto 1/10 2216 as HSA14a and b; HSA15 = raised shiny irregular white/pink mixed, now HSA15a and b 3-2216 restreaked on 1/10 2216

Table 16. Second look on 10/5/12 at H. stipulacea “normal” plates. Inoculum from 1 = undiluted biofilm sample, 2 = 1:5 dilution, and 3 = 1:25 dilution; corresponding re- passaged isolates are described and either re-passaged or stored in glycerol (G) or water. Sample inoculum concentration Qualitative Description of isolates picked earlier and and plate type description of plate action taken beautiful orange pink round convex shiny w/ point, all colonies perfectly stored G/water as HSN26; spread one 1-M13 identical colony on same plate 4 colonies identical stored 3 in G/water as HSN27; restreaked 1-GM13 to HSN26 one on same plate 2 white, 2 coral, 1 black convex shiny 1-anti-2216 spheres, very small restreaking on 2216 as HSN23, 24, 25 113

HSN1 = transparent, dew-like, irregular, shiny, raised - stored G/water; HSN2 = small irregular shiny raised yellow - stored G/water; HSN3 = same as HSN2 - stored G/water; HSN4 = same as HSN1 - stored 1-1/10-2216 G/water HSN5 = brown raised shiny irregular - stored G/water; HSN6 = intense orange raised shiny irregular - stored G/water; HSN7 = raised shiny irregular white and rose, restreaking on 1/10 2216 as HSN7a and b; HSN8 = brown shiny raised irregular - stored G/water; HSN9, 10 same as HSN8 - 1-2216 stored G/water many orangeish stored G/water as HSN30; restreaking one 2-M13 round convex shiny across rest of plate stored G/water as HSN29, restreaking one 2-GM13 identical to HSN30 across rest of plate one coral round 2-anti-2216 convex shiny restreaking on 1/10 2216 as HSN28 HSN21 and 22 - yellow circular raised/flat 2-1/10-2216 shiny - stored G/water as HSN21 HSN19 = yellow-orange raised textured, shiny irregular - stored G/water; HSN20 = white/transparent raised shiny - stored 2-2216 G/water 3 orange raised convex shiny w/ storing 2 colonies G/water as HSN31; 3-M13 point restreaking remaining on same plate fungus-infested except for one picked and streaked on 2216 as HSN32; 3-GM13 orange convex shiny tossed old plate 3-anti-2216 no growth visible HSN11 and 12 - no growth visible; looked at P1 parent plate - nothing unique; tossed 3-1/10-2216 all plates 114

HSN13 = orange-brown shiny raised irregular - stored G/water; HSN14 = dark brown-pink, textured, raised - stored G/water; HSN15 = bright orange raised flat textured - stored G/water; HSN16 = shiny white thick irregular raised - stored G/water; HSN17 = some shiny white-pink irregular raised, some lime-yellow raised/flat textured, restreaking as HSN17a/b; HSN18 = raised shiny dark 3-2216 brown-black, agar-eating - stored G/water

Table 17. Second look on 11/5/12 and 12/5/12 at all plates with inocula originating from flasks; B = broth, S = sludge, sed = sediment; corresponding re-passaged isolates are described and either re-passaged or stored in glycerol (G) or water. Sample inoculum concentration Qualitative description of Description of isolates picked earlier and plate type plate and action taken FHU-M13-B can only see fungus FHU-M13-S can only see fungus possibly beginning of FHU-GM13-B microcolonies saving plate FHU-GM13-S fungus FHU7 = yellow, raised, shiny, FHU-anti- P2: 2 distinct morphologies, irregular; FHU8 = light pink, shiny, 2216-B separate enough to store raised, irregular - stored G/water FHU-anti- FHU9 = same as FHU7; FHU10 = same 2216-S same as broth as FHU8 - stored G/water single colonies easily picked, FHU1 = pink, bumpy, shiny, irregular; many morphologies FHU2 = yellow, shiny, raised, FHU-1/10- represented and re- irregular; FHU3 = same as FHU1/2 but 2216-B passaged light pink FHU4 = pink, convex, shiny, pointy; FHU5 = yellow, shiny, raised, FHU-1/10- irregular; FHU6 = same as FHU4 but 2216-S similar to broth; re-passaged light pink FCS-M13-B fungus FCS-M13-S fungus FCS-GM13-B fungus FCS-GM13-S fungus 115

FCS-anti-2216- P2: pure isolation of yellow- B pink shiny raised irregular stored G/water as FCS1 FCS-anti-2216- S P2: identical to FCS1 stored G/water as FCS2 P2: some white dull raised FCS-1/10- irregular, some shiny yellow restreaking P3 onto 2216 as FCS3 and 2216-B raised irregular FCS4 FCS5 = dull white irregular; FCS6 = yellow raised/flat irregular; FCS7 = pink raised/flat irregular; FCS8 = FCS-1/10- P2: restreaking on 2216 as orange circular w/ dark brown ring 2216-S the following isolates surrounding fungus except for a few FHSA-anti- small pink shiny irregular 2216-ball convex restreaking onto 2216 as FHSA1 note: plate where first on GM13: colonies present but slow- suspected Planctomycetes growing; on 2216: a few more than FHSA-anti- observed; P2 isolates look as GM13 but still slow; on another anti- 2216-S follows 2216: looks same as 2216 FHSA-1/10- 2216-B yellow, shiny, small, convex stored G/water as FHSA2 FHSA-1/10- 2216-S identical to broth stored G/water as FHSA3 many isolated colonies - very very small light pink convex; some have "pooled" FHSN-anti- into large flat/raised stored as FHSN2 and restreaking on 2216-B irregular 2216 FHSN-anti- stored as FHSN1 and restreaking on 2216-S same as broth 2216 FHSN-1/10- beige, flat/raised, irregular, stored G/water as FHSN4 and 2216-B varying sizes restreaking on 2216 FHSN-1/10- stored G/water as FHSN3 and 2216-S same as broth restreaking on 2216 stored some G/water as FHSsed1; P2: very small orange restreaking P2 on 2216; FHSsed2 = microcolonies; not size or also growing poorly, restreaking on FHS-sed-M13 isolated like P1 FHS1 2216 P2: very small FHS-sed- microcolonies; not as storing P1 in G/water as FHSsed3; GM13 healthy looking as P1 restreaking P2 on 2216 116

P2: small orange FHS-sed-anti- microcolonies, similar to 2216-B FHSsed1 restreaking P3 on 2216 as FHSsed9 FHS-sed-anti- 2216-S P2: same as FHSsed9 restreaking on 2216 as FHSsed10 distinct white colonies and orange shiny raised slight restreaking on 2216 as FHSsed6 - FHS-sed-1/10- irregular, also some round orange, FHSsed7 - white, and 2216-B shiny light pink FHSsed8 - light pink FHS-sed-1/10- orange and white irregular restreaking on 2216 as FHSsed4 - 2216-S shiny 'moist' white and FHSsed5 - orange-yellow FCS-sed-anti- P2: small white shiny 2216-B microcolonies restreaking on 2216 as FCSsed1 microcolonies pink, sometimes 'pool,' plate FCS-sed-anti- being overtaken by orange 2216-S fungus storing as FCSsed2 large white shiny raised FCS-sed-1/10- colonies, interspersed with restreaking on 2216 as FCSsed3 - 2216-B convex shiny smaller yellow white, and FCSsed4 -yellow FCS-sed-1/10- 2216-S same as broth FHUsed5 = white shiny microcolonies, restreaking on 2216; FHUsed6 = same as FHUsed5 except pink, restreaking FHU-sed-anti- on 2216 and tossing plate b/c 2216-B overtaken by orange fungus FHUsed7 = same as FHUsed5 except yellow, restreaking on 2216, not enough growth; FHU8 = clear but same as FHUsed7, restreaking on FHU-sed-anti- 2216 and tossing, orange fungus 2216-S taking over FHU-sed-1/10- FHUsed9 = white raised shiny 2216 irregular - stored G/water

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Table 18. Second look on 12/5/12 and 13/5/12 at all plates with inocula originating from sediment; B = broth, S = sludge, sed = sediment, 1 = undiluted original inoculum, 2 = 1:5 diluted original inoculum, 3 = 1:25 diluted original inoculum; corresponding re-passaged isolates are described and either re-passaged or stored in glycerol (G) or water. Sample inoculum concentration Qualitative description of Description of isolates picked earlier and plate type plate and action taken 6 orange-brown convex 1-HS-sed-M13 round restreaking onto 2216 as HSsed1-4 many brown convex shiny 1-HS-sed- circular; one pink convex restreaking as HSsed1-5 and stored GM13 shiny circular G/water; stored G as HSsed1-6 1-HS-sed-anti- 4 convex shiny pink 2216 colonies restreaking on 2216 as HSsed1-1 1-HS-sed- purple agar-eating, circular; stored G/water as HSsed1-7 and 1/10-2216 white dull raised irregular HSsed1-8 orange raised shiny 1-HS-sed- irregular/circular; brown 2216 shiny raised irregular stored G/water as HSsed1-2; HSsed1-3 2-HS-sed-M13 no growth visible 2-HS-sed- 4 orange-brown convex GM13 shiny circular storing G as HSsed2-1 2-HS-sed-anti- 2216 no growth visible 2-HS-sed- HSsed2-2 = purple agar-eating; 1/10-2216 HSsed2-3 = same as HSsed1-8 3-HS-sed-M13 no growth visible 3-HS-sed- 1 orange-brown shiny HSsed3-2 - spreading around same GM13 convex plate 3-HS-sed-anti- 2216 no growth visible tossed 3-HS-sed- 1/10-2216 shiny white raised irregular stored G/water as HSsed3-1 re-passaging 1 on 2216, 1 on M13 as 2 orange-brown shiny CSsed1-7, tossing plate b/c of orange 1-CS-sed-M13 convex fungus one orange-brown convex shiny; agar-eating, orange-brown = CSsed1-3 - 1-CS-sed- crenulated, clearish growth restreaking on remainder of plate, P2 GM13 in middle microcolonies not growing; 118

CSsed1-5 = coral, few raised/flat irregular not doing well - restreaking 1-CS-sed-anti- on 2216; CSsed1-6 = flat/raised pink 2216 irregular - stored G/water 1-CS-sed- orange-white circular shiny 1/10-2216 raised 'moist' stored G/water as CSsed1-4 CSsed1-1 = chunky white raised 'shiny' irregular streaks; CSsed1-2 = same as - 1-CS-sed-2216 1; both stored G/water 2-CS-sed-M13 no growth visible 2-CS-sed- GM13 no growth visible 2-CS-sed-anti- 2216 no growth visible tossed CSsed2-1 = yellow raised shiny irregular/circular - stored G/water; CSsed2-2 = orange dot, created hazy 2-CS-sed- smear around it, not sure if bacterial, 1/10-2216 re-passaging on 2216 1 single orange brown shiny restreaking across rest of plate as 3-CS-sed-M13 convex w/ point CSsed3-4 3-CS-sed- GM13 no growth visible 3-CS-sed-anti- 2216 no growth visible 3-CS-sed- P2: orange-white roundish- 1/10-2216 irregular shiny raised stored G/water as CSsed3-3 CSsed3-1 = orange shiny raised irregular - smells! Not sure if pure, re- passaging; CSsed3-2 = black shiny 3-CS-sed-2216 agar-eating smelly, re-passaging restreaking 2 on 2216 b/c of suspected Planctomycetes as HUsed1- 1-HU-sed- 3 orange-brown colonies, 1, restreaking remaining on new M13 M13 convex shiny w/ point plate b/c this one has fungus 2 orange-brown convex storing in G/water as HUsed1-4; shiny w/ point; 1 orange- restreaking HUsed1-5 on same plate 1-HU-sed- brown flat/raised shiny and keeping P2 for observation b/c GM13 irregular slow-growing 1-HU-sed- anti-2216 no growth visible 119

HUsed1-6 = white raised lumpy circular/irregular shiny jelly-like - stored G/water; HUsed1-7 = same as - 6 but with yellow pigment 1-HU-sed- interspersed throughout, restreaking 1/10-2216 on 2216 for purification HUsed1-2 and -3 are mixed together 1-HU-sed- as orange and white, restreaking for 2216 raised shiny mucus irregular purification 2-HU-sed- M13 no growth visible 2-HU-sed- GM13 no growth visible 2-HU-sed- anti-2216 no growth visible 2-HU-sed- HUsed2-3 = orange-white dew-like 1/10-2216 irregular shiny - stored G/water HUsed2-1 = small brown shiny irregular raised dots; HUsed2-2 = orange-brown mucus shiny large 2-HU-sed- irregular; both grew into each other, 2216 restreaking on separate plates 3-HU-sed- M13 no growth visible 3-HU-sed- GM13 fungus only tossing 3-HU-sed- anti-2216 no growth visible white w/ orange tinge 3-HU-sed- irregular dew-like raised 1/10-2216 shiny stored G/water as HUsed3-1

Table 19. Examination of plates originating from plates inoculated with biofilm, recorded on 13/5/12 and 14/5/12; 1 = undiluted original inoculum, 2 = 1:5 diluted original inoculum, 3 = 1:25 diluted original inoculum; plates were either kept, tossed, or stored in glycerol (G) or water. Isolate number Qualitative description of isolate Action taken HU1a orange-white raised shiny irregular stored G/water transparent orange, 'glows', irregular/round, HU1b raised shiny stored G/water 120

a and b identical; red very irregular flat shiny and producing aqueous gunk; some sections more HU7 pigmented than others stored G/water HU8a purer white irregular shiny/reflective stored G/water HU8b same as HU8a but more yellowish tinge stored G/water brown flat/raised irregular w/ many dots and HU9a pools stored G/water HU9b pink raised shiny irregular stored G/water a and b not distinguishable; dew-like shiny raised HU11 irregular stored G/water HU12a purple raised shiny irregular stored G/water HU12b orange-white raised shiny irregular stored G/water flat/raised, very pigmented deep pink HU13 irregular/circular stored G/water same as HU13 but different shade pink, also has HU14 brown growth with it stored G/water grew well on 2216 but not anti-2216; flat shiny HU31 raised irregular stored G/water moisture present; water/drops forming on anti- HU34 2216 -tossing, not 2216 stored G/water keeping for HU35, 36, 37 originally on GM13; still little growth observation keeping for HU38 originally from M13; little growth observation HU26, 27, 28, 29, 30, 25 all fungus-infested tossed HU41, 42, 43, keeping for 44 slow-growing observation leaving for a few HU19a and b growth visible but not enough to collect days no difference between them; white irregular HU22a and b shiny raised; took over whole plate stored G/water HU23a and b same as HU22 stored G/water HU23c raised shiny clear orange circular stored G/water CS1-anti-2216 fungus tossed CS1-GM13 fungus tossed CS3a white-transparent flat shiny stored G/water CS3b same as CS3a but pink stored G/water CS4a and b watery w/ mucus, dripping, taken over plate stored G/water CS5b orange-white shiny raised irregular stored G/water 121

same as 5b but surrounded by clear-pink slimy CS5a growth stored G/water CS7b orange-white raised shiny irregular stored G/water CS12a white shiny irregular raised stored G/water CS12b same as CS12a except pink stored G/water tossed plate slow-growing pink on anti-2216 - fungus taking after restreaking CS25 over plate on 2216 tossed old plate, restreaked on CS26 same as CS25 but orange 2216 P2 onto GM13 and 2216 completely taken over CS2-GM13 by fungus tossed indistinguishable - shiny irregular white-brown CS24a and b raised/flat stored G/water CS3-M13 no growth since inoculated tossed CS3-GM13 no growth since inoculated tossed streaking remaining colonies on not growing well but 3 shiny pink convex 2216 (from anti- CS28 colonies 2216) very light pink, sheer/transparent, irregular, CS17 shiny, dots/pool stored G/water CS18a same as CS17 but white stored G/water raised pink shiny irregular, contaminated with CS18b white stored G/water brown mucus agar-eating slimy goop with black HSA1 at deepest level stored G/water HSA2 raised shiny irregular orange stored G/water HSA22a and b shiny irregular raised brown-white stored G/water HSA23a orange shiny irregular raised stored G/water HSA23b no growth visible tossed HSA18a white milky shiny mucus watery stored G/water same as 18a but more defined, smaller irregular HSA18b colonies stored G/water HSA20a, b, c orange-white mucus smelly shiny raised irregular stored G/water HSA1-M13 no growth visible since inoculated tossed HSA1-anti- thin film beginning to cover plate but doesn't 2216 seem bacterial tossed 122

keeping for HSA19 slow-growing observation HSA4 yellow, raised, shiny, irregular stored G/water same as HSA4 but different pigmentation, HSA5 slightly more orange stored G/water no pink; only white shiny raised irregular mucus, HSA8a and b smelly stored G/water HSA10 small colonies dull orange, textured stored G/water HSA11a and b orange irregular shiny mucus raised smelly stored G/water HSA3a red shiny irregular- watery stored G/water HSA3b white shiny irregular raised stored G/water HSA25 slow-growing observing HSA2-M13 no growth since inoculation tossed HSA26 P2 onto 2216 (from anti-2216) slow-growing observing HSA27 same as HSA26 observing HSA3-M13 no growth since inoculation tossed HSA28 slow-growing; water beginning to form observing HSA29 P2 beginning to leak water, don't think bacterial tossed HSA12 orange-white raised irregular shiny stored G/water HSA14a yellow shiny raised irregular stored G/water HSA14b same as 14a but white stored G/water HSA15a and b white irregular raised shiny dew-like stored G/water pink and white unable to grow separately; shiny HSA16a and b raised irregular watery stored G/water indistinguishable; 'glowing', orange-clear dots, HSN7a and b raised, shiny stored G/water HSN23, 24, 25 P2 slow-growing, starting to form water observing HSN26 slow-growing observing HSN27 slow-growing observing HSN28, 29, 30 slow-growing observing HSN17a white shiny raised irregular stored G/water HSN17b bright lime irregular -only in streaks- raised shiny stored G/water HSN3-anti- 2216 no growth since inoculated tossed HSN32 since P2 on 2216 from GM13 = slow-growing observing HSN31 slow-growing observing

123

Table 20. Examination of plates originating from plates inoculated from enrichment flasks, recorded on 14/5/12; 1 = undiluted original inoculum, 2 = 1:5 diluted original inoculum, 3 = 1:25 diluted original inoculum; plates were either kept, tossed, or stored in glycerol (G) or water. Isolate number Qualitative description of isolate Action taken FCS5 raised white-orange shiny smelly irregular stored G/water FCS6 white/yellow/clear, dots, shiny and raised stored G/water FCS7 white-pink raised shiny irregular stored G/water FCS8 raised shiny orange-clear tiny dots stored G/water FCS3 white shiny irregular mucus stored G/water FCS4 same as FCS8 except yellow stored G/water light pink-white irregular circular/pools flat FHU1 shiny stored G/water FHU2 same as FHU1 but yellow stored G/water FHU3 same as FCS8 but light pink stored G/water FHU4 same as FHU3 stored G/water FHU5 shiny clear-yellow dots and pools, flat stored G/water FHU6 same as FHU3 stored G/water FHSA1 P2 onto 2216 from anti-2216 slow-growing observing FHSA- suspected all plates -anti-2216, GM13, and 2216 = slow- Planctomycetes growing observing FHSN1 slow-growing observing FHSN2 slow-growing observing white circular shiny raised, stringy, sticks w/ FHSN3 itself and hard to remove from plate stored G/water FHSN4 same as FHSN3 stored G/water FCSsed1-1 slow-growing observing FCSsed1-3 white mucus shiny raised irregular stored G/water FCSsed1-4 small clear-yellow shiny dots, raised stored G/water FHUsed2-5 orange fungus taking over tossed FHUsed2-6 orange fungus taking over tossed FHUsed2-7, 8 death by orange fungus tossed FHSsed1-M13 slow-growing observing FHSsed2-M13 slow-growing observing FHSsed3-GM13 P1 and P2 slow-growing observing white mucus shiny irregular; specks of yellow within zone of antimicrobial activity; avoided FHSsed4 picking them stored G/water FHSsed5 orange shiny circular raised stored G/water FHSsed6 same as FHSsed5 but more intense orange stored G/water 124

textured dull white, very small dotted FHSsed7 colonies stored G/water FHSsed8 small pink clear convex spheres stored G/water FHSsed9, -10 slow-growing observing

Table 21. Examination of plates originating from plates inoculated with sediment, recorded on 15/5/12; the numbers following “sed” (sediment) represent: 1 = undiluted original inoculum, 2 = 1:5 diluted original inoculum, 3 = 1:25 diluted original inoculum; plates were either kept, tossed, or stored in glycerol (G) or water. Isolate number Qualitative description of isolate Action taken CS-sed1-3 slow-growing on GM13 observing CS-sed1-5 no growth visible observing CS-sed1-7 no growth visible observing CS-sed2- GM13 no growth visible observing CS-sed2-M13 no growth visible observing orange shiny irregular raised; grown into agar and weakened it; collected organism along with CS-sed2-2 agar it inhabited stored G/water CS-sed3-1 same as CS-sed2-2 stored G/water CS-sed3-2 same as CS-sed2-2 but uneven pigmentation stored G/water CS-sed3-4 no growth visible on either M13 or 2216 observing HU-sed1-1 observing HU-sed1-2 both shiny raised orange-white, irregular, but -2 and -3 smaller and more reflective w/ thinner streaks stored G/water HU-sed1-5 P1 and P2 not enough growth observing HU-sed1-7 same as HUsed1-3 but yellow stored G/water HU-sed2-1 same as HUsed1-7 but orange stored G/water HU-sed2-2 same as HU-sed2-1 but less orange stored G/water previously anti-2216, now 2216 but still slow- HS-sed1-1 growing observing restreaking on some light orange shiny irregular raised; some 2216 for HS-sed1-2 orange convex shiny smaller dots purification HS-sed1-3 light orange, shiny raised irregular stored G/water HS-sed1-4 slow-growing observing HS-sed1-5, -6 slow-growing observing HS-sed2- GM13 no growth visible observing 125

HS-sed2-M13 no growth visible observing HS-sed3- GM13 no growth visible observing HS-sed3-M13 no growth visible observing

Appendix C: Additional TEM pictures of specimen processed from sample grown on an M13 plate that was streaked from broth enrichment of sediment collected near H. stipulacea

Figure 15. Flagella (fl) and crateriform structures (cr) clearly visible in cell from FHS-sed2 sample 126

Figure 16. Close-up of complex sheathed flagella

127

Figure 17. Specimen displaying polar fibrillar appendages 128

Figure 18. Two cells attached to each other by some unknown mechanism, both displaying polar fibrillar appendages