ASG_Thesis_doublesided

Honours Thesis | A.S. Grobler 1
University of New South Wales
Exploring the ecological role of antibiotic
producing bacteria (APBs) on marine macroalgae
by
Anna Sophia Grobler
A thesis submitted as partial fulfilment of the requirement for the degree of
Bachelor of Science (Honours)
May 2015
Supervised by Dr Suhelen Egan
Centre for Marine Bio-Innovation, School of Biotechnology and Biomolecular Science, UNSW

Honours Thesis | A.S. Grobler i
Originality statement
‘I hereby declare that this submission is my own work and to the best of my knowledge it
contains no materials previously published or written by another person, or substantial
proportions of material which have been accepted for the award of any other degree or
diploma at UNSW or any other educational institution, except where due acknowledgement
is made in the thesis. Any contribution made to the research by others, with whom I have
worked at UNSW or elsewhere, is explicitly acknowledged in the thesis. I also declare that
the intellectual content of this thesis is the product of my own work, except to the extent
that assistance from others in the project's design and conception or in style, presentation
and linguistic expression is acknowledged.’
Signed ……………………………………………..........
Date ……………………………………………..........

ii Honours Thesis | A.S. Grobler

Honours Thesis | A.S. Grobler iii
Acknowledgements
Firstly I would like to thank Dr Suhelen Egan for being a truly wonderful supervisor. Thank
you for your unwavering support over the past year. Without your enthusiasm and guidance
this work would not have been possible. A big thank you also to my mentor, Marwan. With
your help and detailed protocols, I could hit the ground running when I finally received my
DNA sequencing data just over a month ago. Without your assistance, I would undoubtedly
not have been able to analyse these results in time for the submission of my thesis.
Thanks to everyone at BABS, BEES and the CMB, especially the guys in Lab 304 and 306 for
helping me find my feet at UNSW. Becca and Jadi, thanks for assisting me with my fieldwork
and also for the very welcome breaks between long hours in the lab. Special thanks also to
Alex for helping out when I started. Vipra, thanks for your advice and protocols and thanks
Carol for some light entertainment. Mel, thanks for helping me prepare my very first marine
agar plates and showing me how to use ImageJ. Tams, thanks for helping me set up my first
PCR and agarose gel ever and Jun, thank you for showing me how to use the bead beater
and for always helping when I needed to find anything in the lab. Galaxy, you were always
very helpful whenever I needed to order lab supplies - thank you. Meera, thank you for your
advice when I was troubleshooting my PCRs and Leena, thanks for your patient inductions
and for always sending me in the right direction. Nidhi and everyone in Prof Hazel Mitchell’s
lab, thank you for giving me access to the epifluorescence microscope in your lab, and the
interesting chats I had with some of you. Thanks to Jason for helping me with my MiSeq
sample prep, and thanks also to the rest of the guys in Prof Brett Neilan’s and A/Prof Torsten
Thomas’s lab for helping me with odds and ends.
Toni, Tim, James, Das and the guys in the Samuel’s Building, thanks for the quick and always
interesting chats during my breaks, and for some after-hours entertainment. Thanks Tim also
for your feedback on my practice thesis. Esra, thanks for your prompt assistance in getting
my orders placed, and thanks also for the lunch breaks and always lighting up my day with
your beautiful smile. Paris, Pej and Ness, thank you for always sorting me out with anything I
needed in the PC2 lab on level 6. Whether you provided advice, a specific antibiotic or just a
quirky comment and a much appreciated laugh, your kindness will always be remembered.

iv Honours Thesis | A.S. Grobler
I would also like to acknowledge the input of my assessment committee members. Brendan
and Paul, thank you for your helpful, insightful comments and suggestions on my practice
thesis. Thank you both also for making me feel comfortable and confident during the
interview. For financial support, I would like to offer my sincere thanks to UNSW for the
Dean's Honours Relocation Scholarship, as well as to BABS for the BABS Relocation Award
for Honours.
A massive thank you to my family and friends for all your love and support. Thanks for
keeping me up to date with ‘real life’ and helping me to break out of my own head space
once in a while. Mum and Liz, thanks for doing some grocery shopping when I was stuck in
front of the computer or on public transport. Nico, thanks for lending me your mobile phone
when mine broke during the write-up of my thesis. I look forward to spending more time
with you, Edwina and Albert now that it is finally done. Johan, Leanie and Hannah, I cannot
wait to finally come and visit you! Dad, I would have loved to tell you all about my research.
You would have found it just as exciting as I did. I will always miss you.
Lastly to my husband, Johan, thank you for your ongoing support, especially over the past
year. I love you so much and appreciate that you have been my rock in undertaking this
major career change in my life.

Honours Thesis | A.S. Grobler v
Abstract
Ecological interactions between eukaryotes and their associated microbiota are a
fundamental part of life. Such interactions may be positive or negative to the host and
depend on the identity of the microbial partners involved. For example, the natural
antibiotics produced by members of the seaweed microbiome are thought to protect the
host from unwanted colonisation and disease. However, little experimental evidence exists
for the specific role that key microbial players and the antibiotics they produce have in
structuring the microbial diversity on seaweeds and the effect this may have on seaweed
health. Antibiotic producing bacteria (APBs) are hypothesised to benefit their seaweed host
through the production of antimicrobial compounds that structure the functional
composition of the seaweed microbiome. The current study addresses this hypothesis by
investigating whether the APB Pseudoalteromonas tunicata (strain D2) can cause a shift in
the taxonomic composition of epibiotic microbiota associated with the green seaweed Ulva
australis through the production of AlpP, a lysine oxidase with broad antibacterial
properties. The first objective was to develop a protocol whereby P. tunicata could
effectively attach to U. australis surfaces under laboratory conditions. Given that P. tunicata
are known to be ineffective colonisers of mature biofilms, this objective included
investigating methods to disrupt the natural biofilm community without negatively affecting
seaweed health. The results presented here show that both sonication and antibiotics
treatment resulted in a slight, although not statistically significant, decrease in the overall
biofilm coverage on U. australis surfaces. Interestingly, this modest disruption of the
seaweed surface biofilm was sufficient to allow for P. tunicata colonisation of U. australis
surfaces. The second objective was to address the effect of P. tunicata, and in particular the
antibiotic AlpP, on recruitment of the U. australis surface-associated microbial community
(SAMC). U. australis surfaces, pre-colonised with P. tunicata wild type or ΔalpP mutant cells,
were exposed to natural seawater and 16S rRNA gene sequencing was used to investigate
the subsequent recruitment of seawater bacteria to U. australis surfaces. The results
showed a slight shift in the bacterial communities that colonised U. australis surfaces in the
presence of P. tunicata wild type; both when compared to a seawater control, and to the
ΔalpP mutant. In particular, fewer bacteria from the families Flavobacteriaceae and

vi Honours Thesis | A.S. Grobler
Alteromonadaceae colonised U. australis surfaces in the presence of P. tunicata wild type.
Flavobacteriaceae and Alteromonadaceae are known to include algal pathogens and it is
suggested that P. tunicata, through the production of AlpP, inhibited their colonisation of U.
australis surfaces. This study thus demonstrates, for the first time, how P. tunicata and the
antibiotics it produces can play a role in the functional assembly of a microbiome potentially
beneficial to its macroalgal host. These data not only add to the growing understanding of
seaweed-microbe interactions, but the methods developed herein provide the foundation
for future studies aimed at addressing the broader ecological role of APBs in marine systems.

Honours Thesis | A.S. Grobler vii
Table of Contents
Originality statement...................................................................................................................i
Acknowledgements ...................................................................................................................iii
Abstract ......................................................................................................................................v
List of figures .............................................................................................................................. x
List of tables............................................................................................................................... xi
Abbreviations............................................................................................................................ xii
1. Introduction.........................................................................................................................1
1.1. Ecological and economic importance of macroalgae.................................................1
1.2. The importance of the macroalgal microbiome and the holobiont concept.............2
1.3. Macroalgal disease .....................................................................................................4
1.4. Chemical defence in macroalgae................................................................................5
1.5. Antibiotic producing bacteria (APBs) and secondary metabolites............................. 5
1.6. The green macroalga Ulva australis and its microbiome as a model system for
studying the ecological role of antibiotic producing bacteria (APBs) in natural systems .....7
1.7. Study hypothesis.........................................................................................................8
1.8. Study aim ....................................................................................................................9
2. Methods ............................................................................................................................10
2.1. Field collection of U. australis...................................................................................10
2.2. Optimising P. tunicata attachment to U. australis ...................................................10
2.2.1. Comparing the efficiency of methods to disrupt the natural biofilm on U.
australis 10
2.2.2. Determining the effect of biofilm disruption methods on macroalgal health...11
2.2.3. Assessing the ability of P. tunicata to colonise U. australis surfaces.................11
2.3. Assessing the influence of the ABP P. tunicata on subsequent bacterial colonisation
13

viii Honours Thesis | A.S. Grobler
2.3.1. Experimental design and sampling of U. australis microbial community DNA. 13
2.3.2. Extraction of microbial community DNA from the surfaces of U. australis ...... 15
2.3.3. PCR amplification of the 16S rRNA gene............................................................ 15
2.3.4. 16S rRNA gene sequencing using Illumina MiSeq.............................................. 16
2.3.5. Analysis of U. australis microbial community 16S rRNA gene sequences......... 16
3. Results............................................................................................................................... 18
3.1. Efficacy of methods to disrupt the natural biofilm on U. australis...........................18
3.1.2. Fluorescent labelling of bacteria for visualisation after attachment ................ 20
3.1.3. P. tunicata cells were able to attach to sonicated, but not to unsonicated U.
australis surfaces .............................................................................................................. 21
3.2. P. tunicata WT and ΔalpP successfully colonised U. australis surfaces....................22
3.3. Phylogenetic characterisation of U. australis 16S rRNA gene libraries ....................23
3.3.1. Sonication treatment reduced the abundance of Bacteroidetes associated with
U. australis, relative to the abundances of Proteobacteria and Planctomycetes............ 24
3.3.2. Relative abundance of the P. tunicata inoculum and its effect on the richness
and diversity of U. australis microbial communities........................................................ 27
3.3.3. Characterising U. australis communities after the initial 3-hr P. tunicata
colonisation period........................................................................................................... 28
3.3.4. Determining the effect of P. tunicata on the recruitment of microbial taxa from
the natural seawater community to U. australis surfaces............................................... 34
3.3.5. Temporal-driven shifts in community composition........................................... 35
4. Discussion ......................................................................................................................... 36
4.1. P. tunicata was unable to efficiently attach to U. australis without first disrupting
epibiotic biofilms..................................................................................................................36
4.2. Sonication reduced Flavobacteriaceae and Alteromonadaceae, but favoured
Phyllobacteriaceae and Rhodobacteraceae.........................................................................38

Honours Thesis | A.S. Grobler ix
4.3. P. tunicata dominated U. australis surfaces after the initial 3-hr colonisation
period, but did not persist in high abundances after re-introducing the natural seawater
community........................................................................................................................... 39
4.4. P. tunicata, through the production of AlpP, inhibited the colonisation of
Flavobacteriaceae and Alteromonadaceae on U. australis surfaces..................................39
4.5. Summary and future directions................................................................................41
4.6. Concluding remarks ..................................................................................................43
References................................................................................................................................44
Appendix I: Media and buffer solutions...................................................................................53
Appendix II: Example of an agarose gel with PCR products.....................................................57
Appendix III: Summary tables for U. australis community 16S rRNA gene sequencing ..........58

x Honours Thesis | A.S. Grobler
List of figures
Figure 1: The role of microorganisms at different stages in the macroalgal life cycle.............. 3
Figure 2: Habitat and morphology of the green seaweed Ulva australis.................................. 7
Figure 3: The proposed ecological role of antibiotic producing bacteria (APBs) associated
with macroalgal surfaces. .......................................................................................................... 8
Figure 4: Experimental design chart with microbial DNA extraction time points................... 14
Figure 5: Comparing the efficiency of different biofilm disruption methods. ........................ 18
Figure 6: Quantitative assessment of U. australis health after biofilm disruption treatments
and processing (cutting) for the attachment assays................................................................ 19
Figure 7: Attachment of GFP labelled P. tunicata cells to U. australis.................................... 21
Figure 8: Comparing P. tunicata ΔalpP versus WT attachment to U. australis....................... 22
Figure 9: Rarefaction curves showing saturation levels for the detection of bacteria in all the
microbial communities sampled from U. australis and natural seawater. ............................. 23
Figure 10: Multivariate resemblances between microbial communities on sonicated U.
australis compared to the untreated control.......................................................................... 25
Figure 11: Average relative abundances of 97% identity OTUs remaining on sonicated U.
australis surfaces (grey), compared to the no treatment control (white). ............................. 26
Figure 12: Relative abundance (%) of P. tunicata (strain D2) established on U. australis
surfaces. ................................................................................................................................... 28
Figure 13: Richness and diversity of bacterial communities on U. australis surfaces (A)
before; and (B-C) after exposure to natural seawater (NSW) for (B) 24 or (C) 96 hours........ 29
Figure 14: Resemblances between microbial communities associated with U. australis
surfaces (A-C) and natural seawater (NSW). ........................................................................... 31
Figure 15: Average relative abundances of bacterial Classes detected in microbial
communities associated with U. australis surfaces and natural seawater (NSW).................. 32
Figure 16: Heat map showing average relative abundances of bacterial Families detected in
microbial communities associated with U. australis surfaces and natural seawater (NSW).. 33

Honours Thesis | A.S. Grobler xi
List of tables
Table 1: Green fluorescent protein (GFP) expression after 48 hours for P. tunicata ..............20
Table 2: Number of sequences and coverage for 97% identity OTUs......................................58
Table 3: Coverage for 97% identity OTUs subsampled for 1989 sequences............................60

xii Honours Thesis | A.S. Grobler
Abbreviations
APB Antibiotic producing bacterium
CM Chloramphenicol
SAMC Surface-associated microbial community
SSW Sterile seawater
PAM Pulse amplitude modulated
Ƴ Electron transfer quantum yield
PSII Photosystem II
GFP Green fluorescent protein
MB Marine broth (Difco™, Becton Dickinson, USA)
MA Marine agar
O/N Overnight
FOV Field of view
λ Wavelength
OD600 Optical density (absorbance) measured at λ = 600 nm
GCC GFP cell count
TCC Total cell count
bp Base pairs
EPS Extracellular polymeric substance
LPS Lipopolysaccharide
UV Ultraviolet
OTU Operational taxonomic unit

1. Introduction
1.1. Ecological and economic importance of macroalgae
Marine macroalgae - commonly known as seaweeds - are an ancient lineage of
photosynthetic marine metazoans. They include green (Chlorophyta), brown (Pheaophyta)
and red algae (Rhodophyta) (1) and can be distinguished from terrestrial plants by their lack
of vascular structures such as roots (2). Further to their role as primary producers,
macroalgae are important habitat providers on rocky shores (3-7). They shelter a plethora of
invertebrates and fish, including unique animals such as the leafy sea dragon (8,9). In fact,
macroalgae are key ecosystem engineers, and constitute a major structural component of
temperate marine reefs (4,6,10-13). The importance of macroalgae as ecosystem engineers
is further exemplified by the massive decline in fish abundance and diversity which occurred
after large-scale kelp deforestation near California (14). Similarly, invertebrate species
richness off the New Zealand south coast deteriorated after the experimental removal of
two macroalgal species (15). Combined, these data suggest that the removal of habitat-
forming macroalgae will dramatically alter species assemblages in marine systems and may,
in extreme cases, lead to whole ecosystem collapses.
Commercial interest in macroalgae has escalated in recent years and is expected to continue
to rise (16). The demand for edible macroalgae has increased exponentially since the early
1980s (16), and there is a growing interest in using algae for biofuel production (17,18). It is
therefore not surprising to find industry reports stating that the global aquaculture of
macroalgae generates billions of dollars annually and contributes 50% to the total
mariculture industry (19). Marine algae are also exploited in natural products research,
although more recently the search for biologically active compounds from macroalgae has
shifted its focus to their microbial symbionts (20,21). Compared to their macroalgal hosts,
macroalgal-associated microbes are often easier to culture and have shorter generation
times, thus providing a quicker and cheaper avenue for natural products discovery and
production (22,23). Given the economic importance of macroalgae, the results presented in
the current study could provide valuable future applications in the mariculture of seaweed.

2 Honours Thesis | A.S. Grobler
1.2. The importance of the macroalgal microbiome and the holobiont
concept
In aquatic environments, submerged surfaces – whether biotic or inanimate - are quickly
colonised by microorganisms from the water column. Macroalgal surfaces are no exception,
and the density of epiphytic microorganisms associated with macroalgae can vary between
102
to 107
cells.cm-2
, depending on the macroalgal species (24). Macroalgal-associated
microbial communities commonly include representatives from the Proteobacteria,
Bacteroidetes and Planctomycetes (6). Moreover, the microbiomes of macroalgae are
reported to be distinct from the surrounding seawater both in terms of taxonomic diversity
as well as the composition of functional genes (6,25-29).
While a macroalga provides clear benefits to its microbial partners, including a surface for
attachment and a steady supply of nutrients and oxygen, the advantages of these symbiotic
associations to the alga are perhaps less obvious. Microbial biofilms on the surfaces of
seaweeds play an important ecological role in host-environment interactions, analogous to
the function of a second skin (28). The functional outcome of these interactions is, however,
dependent on the identity of the microbial partners present, and may be either beneficial or
detrimental to the host (28,30,31). Macroalgal-associated microbes interact with each other
and their macroalgal host to create a stable system that, under optimal conditions, benefits
them all (6). In fact, the association between macroalgae and their microbiomes has been
proposed to be as important to macroalgal health as the interaction between corals and
their endosymbiotic dinoflagellates, zooxanthellae (6,32). Consequently, the “holobiont”
concept (sensu Rohwer et al. (33)), originally used to describe the tight, obligate relationship
between corals and zooxanthellae, has now been extended to fit the association between
macroalgae and their associated microbes (6,34).
Experimental evidence indicates that macroalgal health critically depends on the close
association with their microbiomes. Macroalgae have complex life cycles, and
microorganisms play an important role in all macroalgal life stages. Microorganisms on the
substrate produce chemical cues that aid in macroalgal “settlement” - the transition of a
motile, planktonic zoospore to its sessile, benthic life stage (35) (Figure 1). The normal
morphological development of many macroalgae depends on bacterially-produced

morphogenesis cues (36) and regulatory factors for cell differentiation and growth (36-38).
For example, without a continuous supply of bacterially synthesised morphogenesis factor
thallusin, abnormal development was observed in the algae Ulva spp. and Monostroma spp.
(36). Macroalgae further depend on microbial partners, such as nitrogen-fixing
cyanobacteria (39), for a constant supply of key nutrients (39,40). Likewise, chemical
defence is costly (41) and it is believed that many seaweed species rely on antibiotic
producing bacteria (APBs) for chemical defence against biofoulers, pathogens, parasites and
grazers, although experimental evidence is still largely lacking in this area. The concept of
chemical defence will be explored in more detail in Section 1.4. Given these important
advantages microbial associates can provide to eukaryotic hosts, a greater understanding of
the formation of beneficial microbiomes in natural systems is required (42).
Figure 1: The role of microorganisms at different stages in the macroalgal life cycle.
Microorganisms on the substrate produce settlement cues, aiding in the transition of a
motile, planktonic zoospore to its sessile, benthic life stage. Many macroalgae further
depend on their microbial partners for morphogenesis cues and growth factors. The surface-
associated microbial community (SAMC) exchange nutrients with their macroalgal host, and
the presence of antibiotic producing bacteria (APBs) is thought to aid in the host’s chemical
defence against biofoulers, pathogens, parasites and grazers.

1.3. Macroalgal disease
As with all eukaryotic organisms, marine macroalgae are susceptible to microbial infection,
disease and decay (16,43-46). Numerous algal disease syndromes have been recorded to
date, including symptoms such as localised thallus bleaching or discolouration, rotting, galls
and lesions (43). Unfortunately, macroalgal disease is increasing in severity and frequency
(10,16,45). This is not surprising, given that scientists have warned against global climate
change and local anthropogenic disturbances, predicting that these disturbances would
increase the occurrence of disease in both terrestrial and marine systems (47).
Over the past four decades, the ocean surface has warmed at an alarming rate (48), and it is
predicted that sea surface temperatures on the East Australian Coast will rise a further 2°C
towards the middle of this century (49). Environmental perturbations such as changes in
temperature and salinity can disrupt the sensitive balance macroalgae have with their
microbiomes (31,50,51) and it has been suggested that the increased incidence of
macroalgal disease under ocean warming conditions may be due to changes in the virulence
of opportunistic pathogens (43).
Whilst some studies argue that macroalgae are relatively resistant to environmental changes
(52,53), the overall increase in macroalgal disease (16) indicates that an improved
understanding of the complex interactions between macroalgae and their associated
microbiota is required. Antibiotic producing bacteria (APB), thought to provide protection
against macroalgal pathogens and aid in the assembly and maintenance of a healthy
microbiome, may be fundamental to macroalgal resilience under rapid environmental
change.

1.4. Chemical defence in macroalgae
Like other sessile marine organisms, macroalgae need mechanisms to defend themselves
against competitors, grazers, parasites, biofoulers, pathogenic invasion and disease.
Deprived of cell-based adaptive immune responses against pathogenic invasion, marine
macroalgae have evolved alternative defence mechanisms such as oxidative bursts
(damaging pathogen DNA), hypersensitive cell death (destroying infected host cells to
restrict the spread of disease) and the production of bioactive compounds (chemical
defences) (43,54-56).
The ability to produce antimicrobial compounds varies between algal species. Rhodophyta
are highly active producers of secondary metabolites against bacterial invasion and
colonisation by biofoulers. For example, the red seaweed Delisea pulchra is known to
produce a range of potent biologically active secondary metabolites (57). Chemical defences
can, however, be energetically costly (41) and as a result, not all macroalgae are rich in
antimicrobial compounds. Chlorophyta such as the edible green seaweed Ulva australis
(commonly known as “sea lettuce”) are, in fact, poor in chemical defence (58). It is
hypothesised that such algae have adapted to rely on antibiotic producing bacteria to
defend them against fouling or pathogenic organisms.
1.5. Antibiotic producing bacteria (APBs) and secondary metabolites
Research on macroalgal-associated microbes carried out thus far has focused mainly on the
discovery of natural bioactive products (e.g. antibiotics) and their potential medical or
commercial application (20,22,58-62). However, less attention has been given to the
ecological role of such bioactive compounds and how they may ultimately affect ecosystem
functioning. The lack of research in this area may largely have been due to difficulties
culturing macroalgal-associated microbes in vitro (63,64). Nevertheless, with technical
advances in culture-independent methods such as high-throughput DNA sequencing and
metabolomics fingerprinting, previously uncultivable organisms can now be studied in detail
(65). Moreover, these next generation “-omics” techniques can be used in manipulative
experiments designed to study how microbial communities change in response to external
stimuli, for example the presence of an antibiotic compound.

As mentioned above (Sections 1.2 to 1.4), both positive and negative interactions exist
between marine macroalgae and their associated microbiota. Compared to their planktonic
counterparts, bacteria in surface-attached biofilms are typically rich in chemical defence
mechanisms (66). While it has been proposed to be an adaptive response compensating for
the loss of mobility to escape predation (59), the abundance of biologically active secondary
metabolites in surface-associated bacteria may also have evolved as a result of increased
competition for space and nutrients.
Epibiotic proteobacteria produce a vast amount of the antimicrobial compounds isolated
from macroalgal surfaces (20,67). For example, the widely distributed Roseobacter clade of
alphaproteobacteria is very abundant in marine systems, often comprising up to 25% of a
bacterial community (68). Within the Roseobacter clade, Phaeobacter species are commonly
associated with green algae (1). Phaeobacter inhibens (formerly Phaeobacter gallaeciensis
(69)) synthesises a potent bioactive antimicrobial compound, tropodithietic acid, that is
proposed to assist their eukaryotic hosts in chemical defence against pathogens and
biofoulers (44,60,70).
Gammaproteobacteria are the most common macroalgal-associated bacterial clade (1).
Pseudoalteromonas species, especially pigmented strains such as Pseudoalteromonas
tunicata, are widely studied gammaproteobacteria due to their ability to produce
antimicrobial compounds (20,71). P. tunicata produces a broad range of bioactive
compounds that substantially inhibit the recruitment and growth of fouling organisms such
as invertebrate larvae, fouling algae, fungi and bacteria (58). One of the many bioactive
compounds produced by P. tunicata is a 190-kDa antibacterial protein, designated AlpP (72).
AlpP inhibits the growth of many medical, terrestrial and marine bacterial isolates, including
both gram-positive and gram-negative bacteria (71,73,74). For example, in vitro experiments
indicated that P. tunicata had inhibitory activity against Staphylococcus aureus, which is a
well-known human pathogen (20). P. tunicata was an affective inhibitor of fungi (75) and
also deterred other fouling organisms such as invertebrate larvae, marine bacteria and algal
spores from settling in petri dishes (60). However P. tunicata had little success at invading
established biofilms on glass or algal surfaces (76).

1.6. The green macroalga Ulva australis and its microbiome as a model
system for studying the ecological role of antibiotic producing
bacteria (APBs) in natural systems
Green macroalgae such as Ulva australis (Phylum Chlorophyta) dominate the intertidal zone
in many temperate marine habitats worldwide (77) (Figure 2). A range of APBs have
previously been isolated from the surface of U. australis (20,60,74,78-80). This study will
focus on one of these isolates, P. tunicata (strain D2), a gammaproteobacterium known to
produce a number of bioactive compounds, including the antibacterial protein AlpP (78). As
introduced in the section above (Section 1.5), previous research has shown that biofilms of
P. tunicata (with specific reference to strain D2) established on plastic surfaces prevented
colonisation by specific bacterial strains, algae, fungi and eukaryotic larvae (60). While these
results strongly suggest the involvement of P. tunicata in macroalgal defence against fouling
organisms, experimental evidence of their respective inhibitory activities directly on U.
australis surfaces is, as yet, not available.
Figure 2: Habitat and morphology of the green seaweed Ulva australis.
(A) The intertidal zone at Clovelly Bay, Sydney, Australia, where specimens were collected
during low tide. (B) The U. australis thallus comprises multiple bright green blades (fronds)
with undulated margins. Stipes are reduced and often completely absent in this species. U.
australis thalli are anchored to rocks in the intertidal zone by a discoid holdfast. Photographs
by An Grobler, 2014.

1.7. Study hypothesis
Given their activity against the colonisation of other microorganisms to macroalgal surfaces
(63), APBs such as P. tunicata have the potential to alter the phylogenetic profiles of
macroalgal-associated microbial assemblages. It is therefore hypothesised that APBs control
ecological interactions within the macroalgal holobiont, thus affecting the functional
properties of these systems to the benefit of their macroalgal host (Figure 3).
Figure 3: The proposed ecological role of antibiotic producing bacteria (APBs) associated
with macroalgal surfaces.
It is predicted that the presence of APBs will alter the surface associated microbial
community (SAMC) to the benefit of the macroalgal host, for example by inhibiting the
colonisation of potential pathogens.

1.8. Study aim
To date, the activities of APBs have only been observed in vitro on plastic and glass surfaces
(58,60,81,82) and virtually nothing is known about the ecological role of APBs associated
directly with the macroalgal surface or their effect on macroalgal health.
Therefore the overall aim of this study is to determine if the model APB P. tunicata, through
the production of the antibacterial protein AlpP, has an effect on the surface-associated
microbial community (SAMC) of U. australis.
This aim will be achieved in the following specific research objectives:
1. Establishing a protocol that will facilitate attachment of P. tunicata on to U. australis
surfaces under laboratory conditions;
2. Determining differences in the phylogenetic composition of the macroalgal
microbiomes that establish on the surface of U. australis in the presence of P.
tunicata wild type (WT) compared to a seawater control; and P. tunicata WT
compared to the ΔalpP strain.

2. Methods
2.1. Field collection of U. australis
U. australis thalli (individuals) were routinely collected from intertidal rock pools at Clovelly
Bay, Sydney, Australia (33.9147o
S, 151.2706o
E, Figure 2 A) during spring and summer
(September 2014 to February 2015). After visible mesograzers (herbivores < 25 mm) had
been removed, macroalgae were kept in aquaria with air bubbling through natural seawater
at 19o
C, with a day/night cycle that consisted of 16 hours light and 8 hours dark.
2.2. Optimising P. tunicata attachment to U. australis
2.2.1. Comparing the efficiency of methods to disrupt the natural biofilm on
U. australis
2.2.1.1. Antibiotic treatment of U. australis samples
Samples were treated according to the protocol originally developed by Rao et al. (74) with
minor modifications as per Dalisay et al. (83). Before antibiotics treatment, 1 cm diameter
discs were excised from U. australis mid-frond sections (Figure 2 B) with a sterile blade and a
glass stencil. Excised U. australis discs were swabbed with NaOCl (0.012%) and incubated in
large, sterile petri dishes at room temperature for 5 min. Discs were then covered in a
broad-spectrum antibiotics mixture that consisted of ampicillin (300 µg.mL-1
), polymyxin B
(30 µg.mL-1
) and gentamicin (60 µg.mL-1
) and incubated for 24 hours at 19o
C. After the
antibiotics treatment, the discs were transferred to fresh petri dishes and incubated in filter-
sterilised (0.22 µm), autoclaved seawater (SSW) for 1 hour at room temperature to remove
residue chemicals.
2.2.1.2. Sonication treatment of U. australis samples
Whole thalli were detached from U. australis holdfasts (Figure 2 B), rinsed with phosphate
buffered saline (PBS, Appendix I) and sonicated for 5 minutes. Sonicated thalli were left to

recover in PBS for 10 minutes after which they were washed three times with PBS and
vortexed between each wash step for 10 seconds. Equal sized discs (1 cm diameter) were
excised from U. australis mid-frond sections (Figure 2 B) with a sterile blade and a glass
stencil. Finally, U. australis discs were placed individually in 1 mL of sterile seawater (SSW) in
mini petri dishes and stored overnight at 19o
C on a shaker (60 RPM).
2.2.2. Determining the effect of biofilm disruption methods on macroalgal health
Macroalgal health after sonication was determined qualitatively - through visual
observations - and quantitatively as described below (Section 2.2.2.1).
2.2.2.1. Pulse amplitude modulated (PAM) fluorometry
The condition of the host was measured before and after the biofilm disruption treatments
using a Mini-PAM (Walz, Germany) according to the manufacturer’s protocol (see Beer et al.
(84) for a simple explanation on how this technique can be applied to Ulva spp). Effective
electron transfer quantum yield (Ƴ) of photosystem II (PSII) was calculated using the
following formula: Ƴ = ΔF/FM, where FM represents the maximal fluorescence and ΔF
represents the variable fluorescence. Mean Ƴ values were calculated based on three
technical replicates for each algal disc.
2.2.3. Assessing the ability of P. tunicata to colonise U. australis surfaces
2.2.3.1. Green fluorescent protein (GFP) labelling of P. tunicata strains
P. tunicata isolates (strain D2) were obtained from the Centre of Marine Bio-Innovation
(CMB) culture collection and revived at 25o
C on half strength marine agar (½ MA, Appendix I)
containing streptomycin (200 µg.mL-1
) for the wild type (WT) and both streptomycin and
kanamycin (85 µg.mL-1
) for the ΔalpP mutant. Strains were labelled with a green fluorescent
protein (GFP) colour tag through conjugation of the plasmid pCJS10G, as described
previously (73). The pCJS10G plasmid contains a gene for chloramphenicol resistance, and
constitutively expresses GFP via activation of the gfpmut3 gene (85). Labelled
transconjugants were maintained on ½ MA containing chloramphenicol (15 µg.mL-1
), plus
kanamycin (85 µg.mL-1
) for the ΔalpP mutant. The stability of GFP expression (excitation λ,

488 nm; emission λ, 510 nm) in newly labelled transconjugants with and without selective
agents in liquid culture after 48 hours was determined by calculating the ratio of viable GFP
expressing cells (GFP cell count, GCC) to the total cell count (TCC) detected using the nucleic
acid stain SYBR® Green I (Molecular Probes, Inc.) (excitation λ, 488 nm (86)).
2.2.3.2. Preparation of bacterial inocula and incubation conditions
P. tunicata WT and ΔalpP inocula were prepared by growing cultures for 12 hours in marine
broth with chloramphenicol (15 µg.mL-1
), plus kanamycin (85 µg.mL-1
) for the ΔalpP mutant.
(25o
C, 180 RPM). Cells were harvested by centrifugation (6,000 x g for 5 min) and washed
twice with SSW before resuspending to an OD600 ~ 0.4. Each replicate seaweed disc (see
Section 2.3.1 and Figure 4 for a detailed description on the experimental design) was
inoculated with 1 mL of bacterial culture at concentrations of approximately 108
cells.mL-1
and incubated at 19o
C for 3 hours whilst shaking gently (60 RPM). Controls were inoculated
with 1 mL of SSW. After the initial 3-hr incubation period, discs from each treatment group
were rinsed in 10 mL SSW to remove unattached bacterial cells and visualised under an
epifluorescence microscope as described below (Section 2.2.3.3).
2.2.3.3. Epifluorescence microscopy
To visualise the amount of natural, live biofilm remaining on U. australis discs after
antibiotics or sonication, biofilms were stained with SYTO® 9 nucleic acid stain and visualised
at an excitation wavelength (λ) of 473 nm using the 40x objective (400x total magnification)
of an epifluorescence microscope (Leica) with a GFP filter (excitation λ, 450-490 nm;
emission λ, 500-550 nm). Attachment of P. tunicata strains to seaweed discs was also
assessed under a GFP filter using the 40x objective. Ten fields of view (FOVs, 224.46 x 167.70
µm each) were captured for each replicate and analysed in ImageJ (87) using the ‘analyse
particles’ function. FOV data were averaged for each replicate and converted to mm2
before
compiling graphs and statistics in Minitab® 17.0 (2013). In all tests for statistical significance,
the significance level was set to α = 0.05.

2.3. Assessing the influence of the ABP P. tunicata on subsequent
bacterial colonisation
2.3.1. Experimental design and sampling of U. australis microbial community DNA
To test the effect P. tunicata has on recruitment of the U. australis microbial community, U.
australis discs - pre-colonised with P. tunicata WT or ΔalpP (Section 2.2.3.2) - were exposed
to natural seawater. Replicate mini petri dishes, each containing 5 mL natural seawater and
a single 1 cm diameter U. australis disc, were given unique identification (ID) numbers to
randomise the treatment groups and incubated at 19o
C (60 RPM) for 24 or 96 hours (Figure
4). ID numbers were used to allocate random spots (using the random list generator on
RANDOM.ORG) for placement of each replicate during incubation. Total microbial
community DNA was collected from U. australis surfaces at five different time points (n = 5
for each treatment group at each time point). Communities were sampled from untreated U.
australis surfaces, after sonication (Section 2.2.1.2), after the 3-hr incubation with P.
tunicata (Section 2.2.3.2), and again after the 24-hr and 96-hr incubation with the natural
seawater community. DNA was also sampled for the natural seawater community (n = 5). All
replicates for which U. australis microbial community DNA was extracted over the course of
the experiment are summarised in Figure 4.

Figure 4: Experimental design chart with microbial DNA extraction time points.
A) Objective 1: Facilitating P. tunicata attachment to U. australis surfaces. After employing
sonication to disrupt the native biofilms on U. australis surfaces, algal surfaces were
A
B

inoculated with P. tunicata wild type (WT), ΔalpP or sterile seawater (SSW) as a control.
Following inoculation, algae were incubated for 3 hours to facilitate P. tunicata colonisation
of U. australis surfaces. B) Objective 2: Determining the effect P. tunicata has on the
assembly of the U. australis microbiome. After the 3-hr colonisation period with P. tunicata,
algal surfaces were incubated with natural seawater for 24 or 96 hours. Total microbial
community DNA was collected from U. australis surfaces at five different time points (n = 5
for each treatment group at each time point). Communities were sampled from untreated U.
australis surfaces, after sonication, after the 3-hr incubation with P. tunicata, and again after
the 24-hr and 96-hr incubation with the natural seawater community. DNA was also sampled
from the natural seawater community (n = 5).
2.3.2. Extraction of microbial community DNA from the surfaces of U. australis
At each DNA sampling time point (Figure 4), replicate macroalgal discs were first rinsed with
SSW to remove any loosely attached cells. Macroalgal surfaces were then thoroughly
swabbed with sterile cotton tips to collect total microbial community DNA from the surfaces.
Microbial biofilm communities (cotton tips) were stored at -20o
C, pending DNA extraction.
Total microbial community DNA was extracted directly from frozen cotton tips using a
PowerBiofilm® DNA Isolation Kit, according to the manufacturer’s protocol (Mo Bio
Laboratories, Carlsbad, CA). Quality and quantity of DNA was determined using a NanoDrop®
ND-1000 spectrophotometer (NanoDrop Technologies, Wilmington, DE).
2.3.3. PCR amplification of the 16S rRNA gene
Fragments of approximately 500 base pairs (bp) were amplified in the polymerase chain
reaction (PCR), using the primer pair 27F/519R to target the V1 to V3 hypervariable regions
of the 16S rRNA gene. The reaction mixture (25 µL total volume per sample) consisted of
EconoTaq® PLUS GREEN 2X Master Mix (Lucigen) (12.5 µL), Ambion® Nuclease-Free Water
(10.5 µL), the primer pair 27F (5’ AGAGTTTGATCMTGGCTCAG 3’) and 519R (5’
GTNTTACNGCGGCKGCTG 3’) (0.5 µL of each; 10 µM) and DNA template (1 µL; products from
Section 2.3.2).

The PCR cycle consisted of initial denaturation at 95o
C (10 min), followed by 35 cycles of
denaturation at 94o
C (30 s), annealing at 55 o
C (10 s) and extension at 72o
C (45 s) with final
extension of 72o
C (10 min). PCR amplicons were visualised on 1% (w/v) agarose gel
containing 0.01% GelRed™ Nucleic Acid Stain (Biotium, USA). Briefly, an aliquot of each PCR
product (5 µL) was electrophoresed with 1× sodium borate buffer (Appendix I) at 70 V (70
min). Amplicon sizes were estimated using a 1-kb DNA ladder (Gene RulerTM) as standard
and photographed under UV illumination (Appendix II) on a Gel-Documentation Unit (Bio-
Rad, Hercules, USA).
2.3.4. 16S rRNA gene sequencing using Illumina MiSeq
Following optimisation of PCR conditions, DNA samples were prepared with indexed primers
(“barcodes”) for Illumina MiSeq with the assistance of Dr Jason Woodhouse, UNSW.
Barcoded PCR products were subsequently sequenced from both ends on the MiSeq Illumina
platform at the Ramaciotti Centre for Gene Function Analysis, UNSW, generating 250 base-
pair (bp) overlapping paired-end reads.
2.3.5. Analysis of U. australis microbial community 16S rRNA gene sequences
Forward and reverse paired-end reads (60 sets) were processed in the programme Mothur
(88) according to the MiSeq standard operating procedure (SOP) (89) (accessed on 2 April
2015). After processing to remove sequencing errors, U. australis microbial community
sequences were aligned to the 16S rRNA molecule using the SILVA database alignment
template (90). Sequences were clustered into operational taxonomic units (OTUs) at a cut-
off of 0.03 and rarefaction curves were created in RStudio (91).
The sequence of the inoculum, P. tunicata (strain D2), was identified using LocalBlast
(version 2.2.30+) (92). After confirming the correct sequence corresponding to P. tunicata
strain D2 in NCBI BLAST (93), the representative OTU (based on 0.03 cut-off) was identified
and its relative abundance was graphed in Minitab (94).
Relatedness within and between U. australis bacterial communities in the different
treatment groups (Figure 4) was explored in PRIMER (95) using multidimensional scaling
(MDS). Permutational multivariate analysis of variance (PERMANOVA) was used to test for

significant differences in U. australis bacterial community composition between samples and
multivariate homogeneity of group dispersions was compared using PERMDISP. Taxonomic
composition was graphed using Minitab (94) and a heat map was assembled in RStudio (91)
using the package ‘heatmap.plus’ (96).

3. Results
3.1. Efficacy of methods to disrupt the natural biofilm on U. australis
Microscopic examination of the algal surface and measurement of total live biofilm coverage
were used to assess the efficacy of biofilm disruption methods via antibiotic treatment or
sonication. Both methods resulted in a slight decrease in biofilm coverage compared to the
untreated controls (Figure 5 A-C). However a one-way ANOVA indicated that these
differences were not statistically significant for either antibiotic (F1, 10 = 0.30; P = 0.597;
Figure 5 D) or sonication treatment (F1, 10 = 2.93; P = 0.118; Figure 5 E). Longer periods of
sonication, including 10 min and 15 min, were not more efficient at disrupting the natural
biofilms on U. australis (F3,20 = 0.30, P = 0.825; Figure 5 E).
Figure 5: Comparing the efficiency of different biofilm disruption methods.
A B C
Livebiofilmremaining(/mm2
)
Livebiofilmremaining(/mm2
)
D E
50 µm 50 µm 50 µm

(A – C) Micrographs represent U. australis surfaces after applying SYTO® 9 nucleic acid stain
to visualise the remaining natural biofilm community per mm2
of algal disc surface. (A)
Untreated control. (B) After antibiotics treatment. (C) After 5 min sonication. (D – E) Graphs
represent the average area of remaining natural biofilm community per mm2
of algal disc
surface after D) antibiotics and E) sonication treatment (n = 6). Bars are one standard error
from the mean. Individual standard deviations were used to calculate the intervals. All
assumptions were met for statistical analysis. Data were approximately normally distributed
and variances were equal.
3.1.1.The effect of biofilm disruption methods on seaweed photosynthetic
efficiency
Photosynthetic efficiency measured with a mini-PAM was used as an indicator of seaweed
health. A one-way ANOVA indicated that antibiotics treatment negatively affected seaweed
photosynthetic efficiency when compared to the control (F1, 10 = 5.18; P = 0.046; Figure 6).
Cutting the seaweed discs did not have a significant effect on photosynthetic efficiency (F1, 10
= 0.83; P = 0.383). Sonication had no effect on photosynthetic efficiency (F1, 10 = 0.55; P =
0.474) relative to the control.
Figure 6: Quantitative assessment of U. australis health after biofilm disruption treatments
and processing (cutting) for the attachment assays.
A B C
Photosyntheticefficiency(ΔF/FMAX
)

Photosynthetic efficiency (ΔF/FMAX), measured using a pulse-amplitude modulated (PAM)
fluorometer, was used as an indicator of seaweed health. Bars are one SE from the mean. A)
Sonication had no effect on photosynthetic efficiency (F1, 10 = 0.55; P = 0.474; One-way
ANOVA). B) Antibiotics treatment negatively affected photosynthetic efficiency when
compared to the control (F1, 10 = 5.18; P = 0.046; One-way ANOVA). Cutting did not have a
significant effect on photosynthetic efficiency (F1, 10 = 0.83; P = 0.383).
3.1.2. Fluorescent labelling of bacteria for visualisation after attachment
To visualise P. tunicata attachment to U. australis discs, strains were labelled with GFP.
Inspection of P. tunicata cells under an epifluorescence microscope indicated that GFP
labelling was successful. Both P. tunicata wild type (WT) and the ΔalpP mutant maintained
GFP expression after 48 hours as indicated by a GCC/TCC ratio of ~1 (Table 1). However the
GCC/TCC ratio was reduced to 0.31 and 0.45 respectively for P. tunicata WT and the ΔalpP
mutant in the absence of antibiotic selection.
Table 1: Green fluorescent protein (GFP) expression after 48 hours for P. tunicata
Strain Growth medium GCC/TCC ratio
Wild type 1/2 MB no antibiotics 0.45
Wild type 1/2 MB SM200 CM15 0.93
ΔalpP mutant 1/2 MB no antibiotics 0.31
ΔalpP mutant 1/2 MB SM200 CM15 0.96
ΔalpP mutant 1/2 MB SM200 CM15 KM85 1.00
GCC = GFP cell count (cells that actively express GFP); TCC = total cell count (this includes all
bacterial cells); SM200 = streptomycin (200 µg.mL-1
); CM15 = chloramphenicol (15 µg.mL-1
);
KM85 = kanamycin (85 µg.mL-1
).

3.1.3. P. tunicata cells were able to attach to sonicated,
but not to unsonicated U. australis surfaces
Previous studies indicate that P. tunicata is unable to efficiently colonise established biofilms
on algal surfaces (76). Therefore the ability of P. tunicata to colonise disrupted, compared to
undisrupted, biofilms on U. australis surfaces was assessed. The use of sonication to disrupt
the natural biofilm present on U. australis had a significant effect on the ability of P. tunicata
to attach (F1, 10 = 23.30; P < 0.001) (Figure 7 C). There were more P. tunicata cells attached to
sonicated U. australis (Figure 7 B) compared to the untreated control (Figure 7 A). In some of
the sonication replicates, microcolonies of P. tunicata had formed after the 3-hr incubation
period, as indicated on Figure 7 B. No microcolonies of P. tunicata were observed on the
untreated control.
Figure 7: Attachment of GFP labelled P. tunicata cells to U. australis.
Micrographs are typical representations of GFP labelled P. tunicata cells on (A) untreated
and (B) sonicated U. australis disc surfaces. Arrows indicate P. tunicata microcolonies. (C)
Data points represent the number of GFP labelled P. tunicata cells that were able to attach
to U. australis discs (n = 6) after 3 hours of incubation at 19o
C. Bars are one standard error
from the mean. Individual standard deviations were used to calculate the intervals.
50 µm
A
50 µm
B
Treatment
SonicatedControl
C
a
b
GFPlabelledcells/mm
2

Variances were unequal. The statistical analysis was therefore performed on log transformed
data. Means that do not share a letter are significantly different (F1, 10 = 23.30; P < 0.001).
3.2. P. tunicata WT and ΔalpP successfully colonised U. australis surfaces
Epifluorescence microscopy was used to confirm that both P. tunicata WT and ΔalpP were
able to colonise sonicated algal surfaces. After three hours of incubation, both strains had
successfully colonised U. australis surfaces. On average, more ΔalpP than WT cells were
attached to U. australis surfaces, however this difference was not statistically significant (F1,8
= 4.27; P = 0.073) (Figure 8).
Figure 8: Comparing P. tunicata ΔalpP versus WT attachment to U. australis.
Micrographs represent (A) P. tunicata ΔalpP and (B) P. tunicata WT cells after the 3-hr
incubation period. Data points represent the number of GFP labelled cells attached per mm2
of U. australis surface (n=5). Bars are one standard error from the mean. Individual standard
deviations were used to calculate the intervals. A one way ANOVA indicated there was no
significant difference in the number of P. tunicata ΔalpP versus WT cells attached to U.
australis (F1,8 = 4.27; P = 0.073).
50 µm
A
50 µm
B
Inoculation
WTΔalpP
C
GFPlabelledcells/mm
2

3.3. Phylogenetic characterisation of U. australis 16S rRNA gene libraries
Analysis of the 16S rRNA gene amplicons across all 60 samples sequenced revealed between
1,989 and 79,939 ~500 bp sequences per sample (Table 2, Appendix III.). Subsampling
reduced the number of sequences to 1,989 per sample (Table 3, Appendix III) for further
analysis. A total of 151,560 unique sequences were identified; 22,549 OTUs at a cut-off of
0.03 (i.e. 97% similarity); and 13,509 at a cut-off of 0.05 (i.e. 95% similarity). Rarefaction data
indicated that the number of new OTUs detected at a cut-off of 0.03 increased steadily, and
sequencing reached saturation after sampling approximately 2,000 sequences (Figure 9).
Figure 9: Rarefaction curves showing saturation levels for the detection of bacteria in all
the microbial communities sampled from U. australis and natural seawater.
Lines indicate the number of OTUs at 97% sequence identity (i.e. 0.03 cut-off) detected
versus the number of sequences sampled from 16S rRNA gene sequences of each microbial
community.

3.3.1. Sonication treatment reduced the abundance of Bacteroidetes associated with
U. australis, relative to the abundances of Proteobacteria and Planctomycetes
To determine the effect of sonication on the taxonomic composition of the U. australis
community, 16S rRNA gene sequences were analysed. Based on OTUs defined at a 97%
identity cut-off, bacterial richness and diversity estimates were, on average, similar on
sonicated U. australis surfaces compared to the untreated control. Sonicated U. australis
communities had an average richness (Chao1) of 417.2, ranging between 236.2 and 598.3;
and an average diversity (inverse Simpson index) of 9.3, ranging between 3.5 and 15.1. One-
way analysis of variance (ANOVA) showed that, at a species cut-off of 0.03, there was no
significant difference in the number of OTUs (F1, 8 = 0.15, P = 0.705), richness (F1 ,8 = 1.04, P =
0.339) or diversity (F1, 8 = 0.01, P = 0.909) of U. australis communities after sonication,
compared to the untreated control.
Multivariate comparison based on Bray Curtis similarity of microbial community structure for
97% identity OTUs revealed slight differences between sonicated U. australis communities
relative to communities in the untreated control (Figure 10). Multi-dimensional scaling
(MDS) showed that, while overall community structure on sonicated U. australis
communities was not significantly different from the untreated control (F1, 5338 = 1.2113; P =
0.3712; PERMANOVA,), replicate U. australis microbial communities within the sonication
treatment group resembled each other more closely, compared to communities in the
untreated control (F1, 8 = 19.329; P = 0.0069; PERMDISP).

Figure 10: Multivariate resemblances between microbial communities on sonicated U.
australis compared to the untreated control.
Multi-dimensional scaling (MDS) based on Bray Curtis similarity of microbial community
structure indicated that sonicated U. australis communities (depicted as squares) were not
significantly different from communities on untreated control replicates (depicted as circles)
(F1, 5338 = 1.2113; P = 0.3712; PERMANOVA). Replicate U. australis microbial communities
resembled each other more closely after sonication relative to the untreated control (F1, 8 =
19.329; P = 0.0069; PERMDISP
Having established general differences between the bacterial communities on sonicated U.
australis compared to the no treatment control, the next step was to investigate which
bacterial taxa contributed to the observed dissimilarities. The relative abundances of 97%
identity OTUs on sonicated algae revealed a slight shift in the ratio of Bacteroidetes,
Proteobacteria and Planctomycetes, with the latter two phyla favoured on sonicated algae,
relative to the no treatment control (Figure 11 A). Bacteroidete abundance on sonicated
surfaces (18.6%) was substantially lower than in the untreated control (27.0%). This was
mainly attributed to a decrease in the abundance of Flavobacteriia (reduced from 15.1% to
6.4%; Figure 11 B) and in particular, Flavobacteriaceae (reduced from 14.4% to 5.6%) after
sonication (Figure 11 C). In contrast, sonication selected for the Saprospiraceaea, a family of
Bacteroidetes within the Sphingobacteria class.

Figure 11: Average relative abundances of 97% identity OTUs remaining on sonicated U.
australis surfaces (grey), compared to the no treatment control (white).
The averages of replicate OTUs (n = 5 for each treatment group) are summarised above,
representing minimum difference (Δ) cut-offs between the treatment groups for (A) Phyla (Δ
A
B
C
PhylumClassFamily

> 0.1%); (B) Classes (Δ > 1%); and (C) Families (Δ > 2%). For simplicity, sequences within each
taxon with negligible differences between the treatment groups (i.e. less than the Δ cut-offs
indicated for A-C) are not displayed.
Proteobacteria after sonication comprised 62.5% of the microbial community, compared to
56.1% on the untreated control (Figure 11 A). Alphaproteobacteria, changing from 35.4% to
45.1% in relative abundance, contributed most to the increase in Proteobacteria on U.
australis after sonication, whilst the relative abundance of Gammaproteobacteria decreased
from 15.1% to 10.2% (Figure 11 B). Phyllobacteriaceae and Rhodobacteraceae contributed
most to the relative increase of Alphaproteobacteria (Figure 11 C). Respectively, the relative
abundance of these two families increased from 1.4% to 4.5% and from 12.6% to 15.3%,
after sonication. The Gammaproteobacteria family, Alteromonadaceae, along with
unclassified Gammaproteobacteria, contributed most to the reduction in the relative
abundance of this Phylum. On untreated U. australis surfaces, Alteromonadaceae had a
relative abundance of 2.8%, compared to only 0.9% after sonication.
3.3.2. Relative abundance of the P. tunicata inoculum and its effect on the richness and
diversity of U. australis microbial communities
Prior to analysing the effect of P. tunicata on U. australis microbial communities, the relative
abundance of the inoculum, P. tunicata (strain D2) was determined. P. tunicata strain D2
was identified within the OTU Otu000001, the most abundant OTU present on U. australis
surfaces after the initial 3-hr incubation, with relative abundances of approximately 23.83%
for the WT and 16.06% for the ΔalpP treatment groups (Figure 12). After exposing the pre-
colonised U. australis surfaces to natural seawater (NSW), the relative abundance of
Otu000001 dropped to 1.00% for the WT and 0.57% for the ΔalpP treatment groups after 24
hours; and to 0.03% and 0.02% for the WT and ΔalpP treatment groups, respectively, after
96 hours. The relative abundance of Otu000001 in NSW was 0.02%. For the SSW control, the
relative abundance of Otu000001 was 0.04% after 24 hours of exposure to NSW, compared
to 0.00% before and 0.00% after 96 hours of exposure to NSW.

Figure 12: Relative abundance (%) of P. tunicata (strain D2) established on U. australis
surfaces.
(A) Before re-introducing the natural seawater (NSW) community. (B) After 24 hours and (C)
96 hours of exposure to NSW.
3.3.3. Characterising U. australis communities after the initial 3-hr P. tunicata
colonisation period
To compare U. australis microbial communities from the different treatment groups after P.
tunicata colonisation, 16S rRNA gene sequencing results were analysed. The results revealed
that, after incubating sonicated U. australis in the presence of P. tunicata WT, microbial
community richness was significantly higher compared to the SSW control (F2,12 = 9.58; P =
0.003; One-way ANOVA; Tukey Pairwise Comparisons; Figure 13 A). This difference could not
be attributed to the production of AlpP, given that there was no significant difference
between the richness of communities in the presence of P. tunicata WT compared to ΔalpP.
There was no difference in the diversity of communities in the presence of P. tunicata WT,
ΔalpP or the SSW control after the initial 3-hr colonisation period (F2, 12 = 0.28; P = 0.761;
One-way ANOVA; Figure 13 A).

Figure 13: Richness and diversity of bacterial communities on U. australis surfaces (A)
before; and (B-C) after exposure to natural seawater (NSW) for (B) 24 or (C) 96 hours.
A) After incubating sonicated U. australis in the presence of P. tunicata WT, microbial
community richness was significantly higher compared to the SSW control, but not when
compared to ΔalpP treatment group (F2,12 = 9.58; P = 0.003; One-way ANOVA; Tukey Pairwise
Comparisons). There was no difference in diversity of these communities (F2, 12 = 0.28; P =
0.761; One-way ANOVA). B) After the 24-hr exposure to NSW, there were no significant
differences in richness (F2, 12 = 0.32; P = 0.734; One-way ANOVA) or diversity (F2, 12 = 1.97; P =
0.182; One-way ANOVA) between the treatments. C) At the 96-hr time point, richness in the
WT treatment varied less than in the SSW control (P = 0.028; Multiple Comparisons) and the
subsequent statistical analysis was performed on log-transformed data. Communities in the
presence of P. tunicata WT were, on average, lower in richness relative to the SSW control,
however this difference was not statistically significant (F2,12 = 0.55; P = 0.588; One-way
ANOVA). There was no significant difference between bacterial diversity in the different
treatment groups at the 96-hr time point (F2, 12 = 0.14; P = 0.871; One-way ANOVA).

Comparison of community composition (presence/absence data based on Bray-Curtis
similarity of 97% identity OTUs) after the 3-hr P. tunicata colonisation period, revealed a
clear difference between communities in the different treatment groups. P. tunicata WT
colonisation had a significant effect on the multivariate resemblance of U. australis microbial
communities relative to the SSW control, but not when compared to the ΔalpP treatment
group (F2, 9948 = 2.4485; P = 0.0267; PERMANOVA; Pair-wise comparisons; Figure 14). In the
presence of P. tunicata WT, replicate U. australis communities resembled each other more
closely than communities in the SSW control (t = 4.9349; P = 0.0081; PERMDISP) and ΔalpP
treatment group (t = 5.4461; P = 0.0098; PERMDISP; Figure 14, circled in orange).
In comparing the relative abundance of OTUs after the 3-hr incubation with P. tunicata,
Bacteroidetes were on average more abundant in the presence of WT (9.9%) compared to
the SSW control (7.0%). Saprospiraceae and Flavobacteriaceae contributed most to this
difference (Figure 16). Saprospiraceae comprised 4.6% of the bacterial community
associated with algal surfaces in the presence of the WT compared to 3.1% in the SSW
control, and Flavobacteriaceae comprised 4.6% of WT community compared to 3.2% in the
SSW control. Similarly, Proteobacteria were more abundant in the presence of WT (80.4%)
and ΔalpP (86.7%), relative to the SSW control (72%). Gammaproteobacteria dominated
microbial communities in the WT (53.6%) and ΔalpP treatments (63.7%), but not in the SSW
control (35.8%). However this difference is in part due to the presence of the inoculum P.
tunicata strain D2. Alphaproteobacteria were more abundant in the SSW control
communities (29.5%) than in WT (23.6%) or ΔalpP communities (19.6%).
The natural seawater microbial community was distinct from U. australis SAMCs in terms of
diversity and overall community structure. U. australis epibiotic communities had lower
species diversity than natural seawater communities (F1, 8 = 5.93; P = 0.04; One-way ANOVA;
Figure 13). Multi-dimensional scaling (MDS) based on Bray Curtis similarity of microbial
community structure indicated that, while OTUs associated with U. australis were distinct
from planktonic seawater communities, replicate U. australis communities resembled each
other (Figure 14).

Figure 14: Resemblances between microbial communities associated with U. australis
surfaces (A-C) and natural seawater (NSW).
Treatments include A) before exposing to NSW; B) after 24 hours and C) after 96 hours
exposure to NSW. The cluster of data points circled in blue (top) represents microbial
communities in NSW. Also note the tight grouping of A-C in the top figure. To more clearly
distinguish between A-C, NSW data points were removed for the bottom figure. Multi-
dimensional scaling (MDS) was based on Bray Curtis similarity. U. australis microbial
communities after the 3-hr incubation period in the presence of P. tunicata WT are circled in
orange (bottom).
NSW
Excluding the NSW data points:

Figure 15: Average relative abundances of bacterial Classes detected in microbial
communities associated with U. australis surfaces and natural seawater (NSW).
The averages of replicate OTUs (n = 5 for each treatment group) from different Classes are
summarised above, representing minimum difference (Δ) cut-offs between the treatment
groups for Classes. Sequences within each Class with negligible differences between the
treatment groups (i.e. Δ < 1%) are not displayed. WT = P. tunicata wild type; ΔalpP = P.
tunicata ΔalpP mutant; SSW control = sterile seawater control. Times in brackets indicate the
different time points at which U. australis microbial communities were sampled during the
current study.

Figure 16: Heat map showing average relative abundances of bacterial Families detected in
microbial communities associated with U. australis surfaces and natural seawater (NSW).
A darker shade indicates a higher relative abundance of a particular taxon. Conversely, a
lighter shade indicates a lower relative abundance. Averages of replicate OTUs (n = 5 for
each treatment group) from different Families were summarised. Sequences within each
Family with negligible differences between the treatment groups (i.e. Δ < 2%) are not
displayed. WT = P. tunicata wild type; ΔalpP = P. tunicata ΔalpP mutant; SSW control =
sterile seawater control. The different time points at which U. australis microbial
communities were sampled during the current study are indicated with brackets.

3.3.4. Determining the effect of P. tunicata on the recruitment of microbial taxa from
the natural seawater community to U. australis surfaces
The effect of P. tunicata on the assembly of the U. australis microbiome was investigated
after re-introducing the natural seawater community to pre-colonised algal surfaces.
Analysis of 97% identity OTU 16S rRNA gene sequences showed that after the 24-hr
exposure to natural seawater, there were no significant differences in richness (F2,12 = 0.32; P
= 0.734; One-way ANOVA) or diversity (F2,12 = 1.97; P = 0.182; One-way ANOVA) between
microbial communities in the different treatment groups (Figure 13 B). Multivariate analysis
of community composition based on Bray-Curtis similarity of 97% identity OTUs also
indicated that, after the 24-hr exposure to natural seawater, there were no statistically
significant differences among, or within, any of the treatment groups (F2, 9921 = 0.5472; P =
0.8788; PERMANOVA, and F2, 9999 = 3.3276; P = 0.066; PERMDISP; Figure 14 B).
Whilst overall, bacterial community structure between the different treatment groups was
not significantly different 24 hours after re-introducing the natural seawater community, a
slight difference in community composition was observed. Bacteroidetes abundance,
although higher for all the treatment groups when compared to before re-introducing the
natural seawater community, was substantially lower in the presence of the WT (17.7%)
relative to the SSW control (22.8%) and ΔalpP treatment groups (22.0%). In particular, there
were less Flavobacteriia associated with U. australis surfaces in the presence of the WT,
compared to the SSW control and the ΔalpP treatment group (Figure 15). The family that
contributed most to this was the Flavobacteriaceae (Figure 16). In the presence of the WT,
notably less Flavobacteriaceae were able to colonise U. australis surfaces (12.6%), compared
to the SSW control (16.9%) and the ΔalpP treatment group (16.5%). There was little
difference in the abundance of Gammaproteobacteria in the three treatment groups (Figure
15). However, differences between families within the Gammaproteobacteria were
observed (Figure 16). In the presence of WT, the abundance of Alteromonadaceae was
substantially lower (14.3%), compared to the SSW control (21.1%) and the ΔalpP treatment
group (19.1%). Conversely, Pseudoalteromonadaceae in the presence of WT was higher
(7.1%), compared to the ΔalpP treatment group (4.1%), and the SSW control (2.0%).

3.3.5. Temporal-driven shifts in community composition
U. australis community richness between the different treatment groups became more
variable over time. After incubating U. australis with the natural seawater community for 96
hours, there was an apparent reduction in the richness of bacterial communities in the
presence of P. tunicata WT, compared to the SSW control, although this was not statistically
significant (F2,12 = 0.55; P = 0.588; One-way ANOVA). Given that the variances were unequal,
with variance in the WT treatment less than in the SSW control (P = 0.028; Multiple
Comparisons), the statistical analysis comparing the means was performed on log-
transformed data. There were no differences in microbial community diversity (F2, 12 = 0.14;
P = 0.871; One-way ANOVA), or between 97% identity OTUs (F2, 9912 = 0.4591; P = 0.8718;
PERMANOVA) at the 96-hr time point (Figure 10).
Compared to the 24-hr time point, the relative abundance of Bacteroidetes associated with
U. australis surfaces at the 96-hr time point had increased substantially for the WT (from
17.7% up to 31.1%) and to a lesser extent for the ΔalpP treatment group (from 22.0% to
30.5%). Conversely, there was a minor increase in Bacteroidetes abundance associated with
the SSW control (from 22.8% to 24.9%). This relative increase in Bacteroidetes abundance at
the 96-hr time point could be attributed to an increase in Flavobacteriia (Figure 15). The
most marked increase in Flavobacteriia abundance was in the WT treatment group (from
13.2% to 23.4%) and could be linked to a rise in Flavobacteriaceae abundance (from 12.6%
to 20.3%).
Between the 24 and 96-hr time points, Proteobacteria abundance declined slightly in the WT
(from 66.8% to 60.7%) and ΔalpP groups (from 68.9% to 60.2%), while increasing in the SSW
control (from 63.5% to 65.7%). This difference was driven largely by the Gammaproteo-
bacteria, while Alphaproteobacteria abundance remained relatively constant from the 24 to
the 96-hr time points (Figure 15). Alteromonadaceae abundance increased slightly for the
WT (from 14.3% to 15.8%) and to a greater extent for the SSW control (from 19.1% to
23.5%), but decreased in the ΔalpP group (from 21.1% to 17.0%).

4. Discussion
Understanding how specific microorganisms can influence the assembly of a beneficial
microbiome is a fundamental question in microbial ecology and evolution (42). APBs,
particularly those isolated from the surfaces of marine eukaryotes such as seaweeds, are
widely studied as a source of novel antimicrobial compounds, but little attention has been
given to the ecological role these organisms have in the natural environment. Whilst the lack
of research in this field may have been due to difficulties encountered using culture-
dependent methods, the rise of culture-independent approaches has opened up new doors
to studying such interactions and gaining insight into the systems APBs form a part of.
4.1. P. tunicata was unable to efficiently attach to
U. australis without first disrupting epibiotic biofilms
Macroalgae are living surfaces with a range of biological, chemical and physical properties
vastly different from non-photosynthetic and inanimate surfaces. The recruitment of
microbiota depends not only on their mechanisms for attachment, but also on other factors
including the availability of oxygen and nutrients, and surface competition through the
production of secondary metabolites (97-99). The observation that P. tunicata was unable to
effectively attach to U. australis surfaces without first disrupting the biofilms via sonication
raised several interesting questions.
Firstly, did sonication select against a small group of OTUs that would, under normal
circumstances, inhibit the attachment of P. tunicata to surfaces with undisrupted biofilms?
Past research demonstrated that P. tunicata was unable to invade mature epiphytic
communities, even when inoculated at high densities (108
cells.mL-1
) (76). Conversely,
another bacterial epiphyte of U. australis, Phaeobacter inhibens (strain 2.10) was able to
invade and colonise the U. australis epiphytic community when inoculated at similar
densities (76). This difference between the abilities of P. tunicata and P. inhibens to invade
the SAMC of U. australis was attributed to the different types of antibiotics produced by
these two strains (76). The 16S rRNA gene sequences of the U. australis SAMC on sonicated,

compared to untreated controls, was also examined in this thesis (see Section 4.2) in an
effort to determine if sonication selected for or against specific taxonomic groups.
Secondly, was there an overall reduction in biofilm biomass after sonication, thereby
creating space on the macroalgal surface for P. tunicata to attach? While the reduction in
total biofilm coverage on algal surfaces after sonication was not statistically significant
(Figure 5), past studies have reported that sonication is effective in the removal of biofilms
(100-102). For example, only brief sonication is required to dislodge bacteria from biofilms
(102), and by combining sonication with vortexing (as in the current study; Section 2.2.1.2)
increases its effectiveness (103). In fact, sonication is the preferred method for the
disaggregation of biofilms from porous surfaces (100,101). Considering the wide use of
sonication to dislodge biofilms from surfaces, together with the observation that P. tunicata
could only efficiently attach to sonicated surfaces compared to the untreated control
(Section 4.1), there may indeed have been a reduction in the overall biofilm biomass after
sonication and additional measurements. Additional methods to quantify biofilm biomass
could be incorporated in future experimental designs.
Finally, could sonication have altered the physical structure of the biofilm extracellular
polymeric substance (EPS) matrix such that it lost its protective ability against antibiotic
invasion? Biofilms are heterogeneous aggregates on surfaces that involves microorganisms
embedded in a thick EPS matrix (104). The EPS matrix makes up about 90% of most biofilms
and provides microbial members that make up the remaining 10% of biofilm with protection
against desiccation, predation and antibiotics exposure (105,106). For example, in mature
biofilms, bacterial cells can be up to a thousand times more tolerant to antibiotics than their
planktonic (suspended) counterparts (106-108). Sonication may have destabilised the EPS
matrix, given that it plays a critical role in bacterial attachment (109), as well as in the
structural integrity and resistance of biofilms (110). Recent advances in the quantification of
EPS in soil biofilms (111) can provide direction for further investigations on the potential of
sonication to alter the composition of the EPS matrix.

4.2. Sonication reduced Flavobacteriaceae and Alteromonadaceae,
but favoured Phyllobacteriaceae and Rhodobacteraceae
U. australis surfaces are typically dominated by Alphaproteobacteria and Bacteroidetes (80).
Sonication - used in the current study to disrupt the biofilms naturally present on U. australis
surfaces - showed strong selective activity against Flavobacteriaceae (Figure 16), a
Bacteroidetes family that is usually abundant on U. australis surfaces (80). Members of the
Flavobacteriaceae are known to cause disease in fish, and are a major threat to the
aquaculture of salmon (112,113). Some Flavobacteriaceae strains produce secondary
metabolites that may inhibit the growth or recruitment of other bacteria (114). For example,
the flavobacterium Zobellia galactanivorans, commonly associated with marine macroalgae,
produces bioactive compounds through a vanadium iodoperoxidase (114). Gammaproteo-
bacteria, especially within the Alteromonadaceae, a family with members known to produce
antimicrobial compounds (115), were also selected against during sonication (Figure 16).
Considering their potential to produce antimicrobial compounds, bacteria from the
Alteromonadaceae and Flavobacteriaceae families have the potential of inhibiting P.
tunicata recruitment to U. australis surfaces. It is therefore possible that sonication reduced
the amount of biofilm members able to produce inhibitory compounds against other
bacterial strains, such that P. tunicata were able to colonise U. australis surfaces.
Specific families within the Alphaproteobacteria demonstrated a higher degree of resistance
to sonication, the Phyllobacteriaceae and Rhodobacteraceae. Alphaproteobacteria share a
long evolutionary history with higher eukaryotes (116,117) and are common symbionts
associated with plants and animals (118). Subsequently, this group of bacteria have evolved
mechanisms to colonise, and remain in close association with, their host. Rhizobia, for
example, are nitrogen-fixing endosymbionts of legumes that infect their hosts via adhesins
and cellulose fibrils (119). In marine systems, members of the Rhodobacteraceae are well
adapted to life on eukaryotic host surfaces, producing extracellular polysaccharides that
contribute to their ‘stickiness’ (120). P. inhibens is a member of the Rhodobacteraceae family
and, considering the roles it may have in algal defence (introduced in Section 1.5) and
biofilm formation, would be an interesting species to include in future studies regarding the
ecological role of APBs associated with U. australis (discussed further in Section 4.5).

4.3. P. tunicata dominated U. australis surfaces after the initial
3-hr colonisation period, but did not persist in high abundances after
re-introducing the natural seawater community
Although P. tunicata were able to colonise and form dense films on sonicated U. australis
surfaces, these - apparently well-established biofilms of P. tunicata mixed with the taxa that
remained after sonication - were unable to persist over the course of the experiment
(Section 3.3.2). This raises a number of new questions that can be addressed in future
studies. First, was this failure of P. tunicata to remain associated with U. australis surfaces
due to the recovery of a specific taxon’s competitive ability to exclude P. tunicata from the
U. australis microbial assemblage? Second, could the inability of P. tunicata to persist be due
to the aggressive colonisation by another taxon from the natural seawater community?
Third, did the physical conditions on the host surface change and become unfavourable for
P. tunicata to persist, leading through a natural succession of bacteria, ultimately favouring
organic decomposers? Despite the significant reduction in abundance of P. tunicata over
time it was still present on U. australis surfaces - albeit at a lower relative abundance - after
24 hours of exposure to the natural seawater community and therefore the effect of P.
tunicata on the SAMC could be explored further.
4.4. P. tunicata, through the production of AlpP, inhibited the
colonisation of Flavobacteriaceae and Alteromonadaceae
on U. australis surfaces
In addressing the main thesis hypothesis, it was clear that members of the Flavobacteriaceae
and Alteromonadaceae were less successful at colonising U. australis surfaces in the
presence of P. tunicata WT relative to the both the P. tunicata ΔalpP treatment and the SSW
control (Section 3.3.4). Macroalgal pathogens have been reported for both these families,
specifically from the genera Alteromonas and Flavobacterium (121-124). For example,
Alteromonas species induced lesions in previously healthy thalli of the brown macroalga,
Laminaria religiosa (121), and caused gametophyte infection in L. japonica (122). Laminaria
is an economically important genus of seaweed that is extensively cultivated for the
production of alginate - a thickening agent widely used, for example, in ice cream (125).

Flavobacterium sp. was the causative agent of the so-called “anaaki disease” (a series of
pinhole-like perforations) in the red macroalga, Porphyra yezoensis (124). Porphyra species,
popularly known in Japanese cuisine as “nori”, are also widely cultivated and thus provides
another example of economically important seaweed genus (124).
Previously, comprehensive studies on the diversity of U. australis epibiotic bacterial
communities demonstrated that typical recruits to these hosts include (among others)
Flavobacteriaceae and Alteromonadaceae, with higher abundances on U. australis surfaces
compared to the surrounding seawater (61,80,126,127). In light of these previous studies, it
stands to reason that high relative abundances of these taxa, if present in the natural
seawater community, would recruit to U. australis surfaces. The low relative abundance of
Flavobacteriaceae able to colonise in presence of the WT compared to the ΔalpP treatment
group suggests that P. tunicata, through the production of AlpP, can competitively inhibit
specific members of the Flavobacteriaceae from colonising U. australis surfaces. P. tunicata
may also produce bioactives against Alteromonadaceae, given the low relative abundance of
this family in both the WT and ΔalpP treatment groups after the initial 3-hr P. tunicata
colonisation period compared to the SSW control. Moreover, even with a dramatic decline in
the inoculum at the 96-hr time point, Alteromonadaceae numbers in the WT and ΔalpP
treatment groups remained significantly lower than in the SSW control (Figure 16).
Given the diversity of antimicrobial compounds produced by P. tunicata (58,59,75,128,129)
it is possible that, in addition to AlpP, P. tunicata produces other bioactives that target
specific strains within the Alteromonadaceae. For example, P. tunicata also produces
violacein, a purple pigment known to have broad antibacterial activity (130,131), as well as
YP1, a yellow tambjamine pigment (79). Determining the specificity of bioactives such as
violacein and tambjamine against particular bacterial strains could provide an interesting
topic for further investigation, particularly in the discovery and design of novel antibiotics.
Finally, whether the reduction in Flavobacteriaceae and Alteromonadaceae abundance in
the presence of P. tunicata was due to the inhibition of growth of these families already
present on U. australis surfaces, or due to the inhibition of recruitment from the natural
seawater community, remains to be explored in future research.

4.5. Summary and future directions
In summary this thesis demonstrated that, while P. tunicata are inefficient colonisers of U.
australis surfaces with established biofilms, sonication can disrupt these biofilms to facilitate
P. tunicata attachment. In addition, it showed that P. tunicata, likely through the production
of AlpP, inhibited the colonisation of members of the Flavobacteriaceae and
Alteromonadaceae on U. australis surfaces. These data not only add to the growing
understanding of seaweed-microbe interactions (3,6,25), but the methods developed here
provide the foundation for future studies aimed at addressing the broader ecological role of
APBs in marine systems.
Whilst the use of Illumina 16S rRNA gene amplicon sequencing provides a good baseline for
community changes based on phylogenetic identification of bacterial species, the relatively
short reads (< 500 bp) means that in many cases it is not possible to obtain high-resolution
taxonomic identification (i.e. to genus or species level). Alternatives to the 27F/519R primer
combination could also be explored to capture greater 16S rRNA gene sequence diversity,
for example the primer set 356F/1064R has been proposed to increase coverage for the
detection of bacterial diversity by 50% (132). Moreover, phylogenetic analysis does not
provide details on the functional genes present in a microbial community. Given than
microbial community assembly has been linked to function, not taxon (127), future work
should consider metagenomic sequencing (65,133) to supplement these data. Previously,
metagenomic approaches have been applied successfully to study the SAMC of U. australis
(20,65,80,127,133). For example by using a functional genomics approach, an antifungal
biosynthetic gene cluster was identified in the P. tunicata genome (133).
The current study forms part of a larger, more comprehensive study on the ecological role of
APBs on marine macroalgae. As such, it is intended to provide a foundation for future work
in this area. It is recommended that the future studies include a more comprehensive
approach in assessing the health of U. australis. In addition to the use of PAM fluorometry to
measure photosynthetic efficiency, other measurements of algal health can be considered.
For example, cell viability can be assessed by staining algal cells with Evans Blue Dye, a non-
toxic dye that penetrates non-viable cells (134,135). Ideally, this research should be carried
out on whole thalli over a longer time to assess health more accurately. By scaling up the

techniques developed here, they can be applied to whole U. australis kept in suitable
mesocosm style aquaria - designed for this purpose - at the Sydney Institute for Marine
Science (SIMS).
In addition to bacteria, microscopic eukaryotes such as fungi are also important components
of microbial communities, and can dramatically alter microbial interactions (136). While the
current study focused on characterising how bacterial partners of the microbial community
respond to the presence of an APB, the microbial community DNA extracted here is a
valuable resource intended for further investigations to generate a more comprehensive
understanding of the effect of P. tunicata on the SAMC of U. australis. Such future work can
supplement the 16S rRNA sequencing data generated in the current study by using 18S rRNA
gene sequencing to characterise the eukaryotic partners of this community (137).
Determining the ability of P. tunicata to persist for longer on U. australis surfaces may also
provide an interesting topic for further investigation. For example, longer incubation periods
may be required to facilitate P. tunicata establishment on U. australis surfaces. Moreover
cooperative biofilm formation between P. tunicata and other marine strains has been
suggested previously (74) and based on this, experiments could be designed to test the
ability of P. tunicata to colonise U. australis surfaces in the presence of specific bacterial
strains. Given that many environmental bacteria are thought to be better adapted to
dispersal versus the biofilm formation (138), the possibility that P. tunicata does not
maintain itself over long periods also needs further exploration. The ecological role of other
APBs previously isolated from U. australis surfaces can also be included in future work. For
example, does P. inhibens (introduced in Section 1.5) through the production of bioactives
such as tropodithietic acid, structure the SAMC of U. australis to the benefit of the algal
host?

4.6. Concluding remarks
To date, most of the research regarding APBs isolated from marine eukaryotes has focused
on their antimicrobial activities, with the aim of discovering novel antibiotics. However, the
ecological roles these microorganisms and their antimicrobials have within their respective
host-associated micro-niches, particularly with respect to microbiome assembly, are still
poorly understood. Microbiomes are known to promote a range of eukaryotic host
functions. Hence, elucidating the mechanisms underlying the assembly of microbiomes
beneficial to eukaryotic hosts has become one of the leading questions in biology, attracting
researchers from diverse fields. In aquatic biology, studies in molecular ecology have
demonstrated that fish microbiomes can assist their hosts in acclimatisation to rapid
environmental changes (139). Shifts in the microbial communities of corals, with particular
emphasis on the absence of beneficial members of the microbiome, are implicated in coral
disease (140). There are numerous terrestrial examples investigating the functional and
phylogenetic assembly of microbiomes, such as studies on the assembly of healthy root-
associated rice microbiomes (141). The role native microbiota of humans play in normal
infant development (142) and health maintenance (143) has also received increased
attention in recent years. Interestingly, many of these studies draw on ecological theory
when attempting to predict the role of key players in microbiome assembly. Thus, in
exploring the ecological role of APBs on seaweed hosts, this study contributes to the general
understanding of APBs in their natural environment. Combined with the results from other
studies, it may ultimately allow microbiologists to address fundamental questions in the
ecology and evolution of microorganisms and in particular, how eukaryotes, whether they
are macroalgae, corals or even humans, assemble microbiomes that are beneficial to their
health.

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