UNIVERSITY OF COPENHAGEN
FACULTY OF SCIENCE
DANISH ARCHAEA CENTRE
PhD thesis
Kristine Buch Uldahl
Hyperthermophilic archaeal viruses as
novel nanoplatforms
Academic supervisor: Xu Peng
November 2015
UNIVERSITY OF COPENHAGEN
FACULTY OF SCIENCE
DANISH ARCHAEA CENTRE
PhD thesis
Kristine Buch Uldahl
Hyperthermophilic archaeal viruses as novel
nanoplatforms
Academic supervisor: Xu Peng
November 2015
Institutnavn:
Biologisk Institut
Name of department:
Department of Biology
Section:
Functional Genomics
Author:
Kristine Buch Uldahl
Titel:
Hypertermofile arkæavirus som nye nanoplatforme
Title / Subtitle:
Hyperthermophilic archaeal viruses as novel nanoplatforms
Subject description:
This thesis aims at evaluating archaeal viruses as novel nanoplatforms.
The focus will be on investigating the hyperthermophilic archaeal virus,
SMV1, to gain insights into the viral life-cycle and to provide a strong
knowledge base for developing SMV1 into a nanovector platform.
Main supervisor:
Associate Professor Xu Peng
Co-supervisor:
Professor Moein Moghimi
Submitted:
November 2015
Type:
PhD thesis
Cover:
Top: Kristine Uldahl, sampling trip Yellowstone National Park, Left:
TEM image of Sulfolobus monocaudavirus 1, Bottom: Morning Glory
Hot spring, Yellowstone National Park
Preface
This thesis is the product of a three-year PhD project at the Faculty of Science, University of
Copenhagen, based at the Danish Archaea Centre, Department of Biology. The thesis has been
supervised by associate Professor Xu Peng with heavy involvement from co-supervisor Professor
Moein Moghimi (Centre for Pharmaceutical Nanotechnology and Nanotoxiocology (CPNN),
University of Copenhagen). Further guidance, collaboration, and advice were received in relation to
specific chapters from Mark J. Young and Seth T. Walk.
The thesis consists of two parts. The first part is a synopsis which gives an overview of the
background and objectives of the thesis, summarizes and discusses the main findings, and outlines
some perspectives for future research. The second part consists of 4 manuscripts, written as
scientific papers, which comprise the core of the PhD project.
During my PhD I had the pleasure of spending approximately half a year in the lab of Mark J.
Young at Montana State University. Further, I was invited by Dr. Brent Peyton (Head of the
Thermal Biology Institute, Montana State University) to participate in the annual sampling trip to
the Hot Springs at Heart Lake, Yellowstone National Park. The stay at Montana State University
was partially funded by Knud Højgaards Fond and Oticon Fonden.
Besides the work contained in the chapters I have also assisted with teaching the course, Archaea
Biology, for two consecutive years and supervised two bachelor students with laboratory
experiments and troubleshooting.
Kristine Buch Uldahl
Copenhagen, Denmark
November 2015
Table of contents
1. Preface
v
2. Table of Contents
vii
3. Summary
1
4. Sammenfatning
3
5. Acknowledgements
5
6. List of manuscripts
6
7. Synopsis
7.1
Introduction
7
7.2
Aims of the Thesis
8
7.3
Archaea: the new branch on the tree
9
7.4
The wondrous world of archaeal viruses
16
7.5
When viruses become useful
25
7.6
Design strategies of viral nanoplatforms
27
7.7
From engineering to applications in medicine
32
7.8
Summary of results
33
7.9
Outlook
38
7.10 References
40
8.
Chapter I
45
9.
Chapter II
69
10. Chapter III
97
11. Chapter IV
127
Summary
Viruses are the most abundant biological entities on earth, and with an estimated 1031 virus-like
particles in the biosphere, viruses are virtually everywhere. Traditionally, the study of viruses has
focused on their roles as infectious agents. However, over the last decades with the development of
a broad range of genetic and chemical engineering methods, viral research has expanded. Viruses
are now emerging as nanoplatforms with applications in materials science and medicine. A great
challenge in biomedicine is the targeting of therapeutics to specific locations in the body in order to
increase therapeutic benefit and minimize adverse effects. Virus-based nanoplatforms take
advantage of the natural circulatory and targeting properties of viruses, to design therapeutics that
specifically target tissues of interest in vivo. Plant-based viruses and bacteriophages are typically
considered safer nanoplatforms than mammalian viruses because they cannot proliferate in humans
and hence are less likely to trigger adverse effects. Another group of viruses that fits this criterion is
archaeal viruses yet their potential remains untapped. As a group, archaeal viruses offer distinct
advantages such as unique morphotypes and inherent stability under extreme conditions. This thesis
presents the first in depth investigation of any archaeal virus, SMV1, as a potential nanoplatform for
applications in nanomedicine.
In order to provide a strong foundation for downstream experiments and future applications,
Chapter I presents an in depth investigation of the hyperthermophilic archaeal virus SMV.
Decisive steps in the viral life-cycle are studied with focus on the early stages of infection. TEM
observations suggest that SMV1 virions enter into host cells via a fusion entry mechanism,
involving three distinct stages; attachment, alignment, and fusion. Upon infection, the intracellular
replication cycle lasts 8 h at which point the virus particles are released as spindle-shaped tailless
particles. Chapter II builds on the replication and purification methods in Chapter I to study the
interaction between the two hyperthermophilic archaeal viruses, SMV1 and SSV2 and cells of
human origin. This chapter provides the first results demonstrating that archaeal viruses can be
taken up and internalized by human cells, thus indicating a potential as intracellular delivery agents.
Chapter III investigates SMV1 particles as potential nanocarriers targeting the gut microbiome.
Stability experiments proved SMV1 to be highly stable in both simulated conditions of the human
gastrointesinal tract (in vitro) and when passaged orally in mice (in vivo). In general, high doses of
SMV1 elicited a nearly undetectable murine inflammatory response and challenged mice showed no
1
observable signs of pain or distress. The stability of SMV1 was compared to that of the traditionally
used Inovirus, M13KE. SMV1 outperformed this state-of-the-art vector as measured by in vitro and
in vivo survival. Chapter III provides strong evidence that SMV1 in particular and archaeal viruses
in general have intrinsically favorable in vivo characteristics for bioengineering applications, such
as drug delivery in the gastrointestinal tract. Chapter IV presents an overview of all known
archaeal viruses and discusses the application potential of archaeal viruses.
2
Sammenfatning
Vira er de mest udbredte biologiske enheder på jorden. Vira er næsten overalt. Det er anslået at der
er 1031 viruslignende partikler i biosfæren. Traditionelt set har studiet af virus fokuseret på deres
rolle som sygdomsfremkaldende agenter. Men i de seneste årtier med udviklingen af en bred vifte af
genetiske og kemiske teknikker har virusforskning udvidet sig. Vira finder nu anvendelse som
organiske nanoplatforme indenfor materialevidenskab og medicin. En stor udfordring i biomedicin
er målretningen af lægemidler til bestemte steder i kroppen for at øge terapeutisk fordel og
minimere bivirkninger. Virus-baserede nanoplatforme drager fordel af naturlige virusegenskaber i
form af nanostørrelse og symmetri, til at designe lægemidler, der er målrettet specifikt væv i
kroppen. Vira der inficerer planter og bakterier, bliver typisk betragtet som ”mere sikre” i forhold til
eukaryotiske vira, da de ikke kan reproducere sig i mennesker og dermed er mindre tilbøjelige til at
forårsage bivirkninger. En anden gruppe af vira der passer til dette kriterium, er arkæavira. Deres
potentiale er endnu ikke blevet udnyttet. Som gruppe tilbyder arkæavira flere fordele såsom unikke
morfologiske former og naturlig stabilitet under ekstreme forhold. Denne PhD afhandling
repræsenterer den første dybtgående evaluering af en arkæavirus, SMV1, som en potentiel
nanoplatform til applikationer indenfor nanomedicin.
For at give et stærkt fundament for efterfølgende eksperimenter og fremtidige applikationer,
præsenterer Kapitel I en dybtgående undersøgelse af den hypertermofile arkæavirus SMV1.
Afgørende faser i den virale livscyklus studeres med fokus på de tidligste faser af infektionen. TEM
observationer tyder på at SMV1 partikler trænger ind i værtens celler via en fusionsmekanisme, der
involverer tre forskellige stadier; vedhæftning, positionering, og fusion. Efter infektion varer den
intracellulære replikationscyklus 8 timer, på hvilket tidspunkt viruspartiklerne frigives som haleløse
partikler. Kapitel II bygger på oprensningsmetoder fra Kapitel I for at studere samspillet mellem
de to hypertermofile arkæavira, SMV1 og SSV2 og celler af menneskelig oprindelse. Dette kapitel
indeholder de første resultater der viser at arkæavira kan optages og internaliseres af humane celler,
hvilket indikerer et potentiale som intracellulære leveringsagenter. Kapitel III undersøger SMV1
partikler som potentielle nanocarriers rettet mod tarmmikrobiomet. Stabilitetsforsøg viste at SMV1
partikler er særdeles stabile i både simuleret tarmmiljø (in vitro), og under tarmpassage hos en mus
(in vivo). Generelt set gav høje doser af SMV1 et ikke målbart murin inflammatorisk respons og
udfordrede mus viste ingen observerbare tegn på smerte eller stress. Stabiliteten af SMV1 blev
3
sammenlignet med den traditionelt anvendte Inovirus vektor, M13KE. SMV1 udkonkurrerede
denne state-of-the-art vektor både målt ved in vitro og in vivo. Kapitel III giver stærke beviser for,
at SMV1 især og arkæavira generelt har gunstige in vivo egenskaber for bionanoteknologiske
applikationer. Kapitel IV indeholder en oversigt over alle kendte arkæavira og diskuterer
anvendelsespotentialet for arkæavira i almindelighed.
4
Acknowledgements
This PhD could not have been carried out – and would not have been as good an experience –
without the involvement of a larger number of people. First and foremost I would like to thank all
colleagues, family and friends for the support and interest during the past three years. A big thank
you goes out to everybody at the Danish Archaea Centre (DAC) and many good colleagues and
friends at the Department of Biology for providing an inspiring and enjoyable work environment
over the past years. Furthermore, I need to thank the Thermal Biology Institute at Montana State
University for hosting me during my half year research stay; in particular everybody in the Young
lab who made my stay in Bozeman truly amazing. Thank you to Dr. Brent Peyton and the Heart
Lake crew for a marvellous trip into Yellowstone (2014), a truly amazing experience.
This PhD is to a high degree a product of two external research collaborations. I want to thank Prof.
Mark J. Young and Prof. Seth T. Walk for collaborations introducing “my virus” to mice and for
their scientific encouragement at a much-needed time. I need to thank Prof. Moein Moghimi and
PhD LinPing Wu for their enthusiasm and ideas without which my project would not have been the
same.
Almost last, but certainly not least, I would like to thank my supervisor, Prof. Xu Peng. Thank you
for the past three years and for the opportunity for me to develop as a person and scientist. I have
valued our scientific discussions and have learned a lot about broader aspects of science and critical
scientific thinking from you.
I feel fortunate to have a wonderful family and amazing friends. I cannot express in words the
importance of knowing that you are all there when I need it. I extend my gratitude to my mum,
Allan and Thomas for their unconditional love and support. A gigantic thanks to Bo for
accompanying me to Yellowstone and beyond, you have been my greatest support and fill my life
with love.
5
List of manuscripts
I.
Uldahl, K. B., Jensen, S. B., Bhoobalan, Y., and Peng, X. (2015) First insights into the entry
mechanism of a spindle-shaped extremophilic archaeal virus and its interaction with
Sulfolobus host, manuscript in preparation
II.
Uldahl, K. B., Wu, L., Hall, A., Peng, X., and Moghimi, M. (2015) Recognition of
extremophilic archaeal viruses by eukaryotic cells; an emerging nanoplatform from the third
domain of life, submitted Biomaterials
III.
Uldahl, K. B., Walk, S. T., Olshefsky, S. C., Peng, X., and Young, M. J. (2015) SMV1, an
extremely stable thermophilic platform for nanoparticle trafficking in the mammalian GI
tract, submitted PlosOne
IV.
Uldahl, K. B. and Peng, X. (2015) Review: Emerging nanoplatforms based on extremophilic
archaeal viruses, manuscript in preparation
6
Synopsis
Hyperthermophilic archaeal viruses as novel nanoplatforms
1
Introduction
The Archaea evolved as one of the three primary lineages several billion years ago, but the Archaea
were formally proposed as the third domain of life only 25 years ago (1). In this short period of
time, scientists have identified many unique characteristics about the Archaea – from distinctive
cellular pathways to their abundance and critical function in diverse natural environments (2). And
not least, the discovery of the Archaea opened the door to the amazing world of archaeal viruses.
Recent research has revealed that DNA viruses of the Archaea have highly diverse and often
exceptionally complex morphotypes. Moreover, studies on viral life-cycles have shown some
exceptional mechanisms including; viral release through pyramid-like structures, extracellular
morphological development of virions, and virus-induced dormancy of host cells (3, 4). All known
archaeal viruses have been isolated from extreme environments, primarily from terrestrial hot
springs and hypersaline lakes. Their habitats are often characterized by more than one extreme and
fluctuations are common (5, 6). Just as their hosts, archaeal viruses have had to adapt to withstand
these extreme environmental conditions, making them extremely stable nanoparticles.
Initially the unique morphotypes of archaeal viruses stole the spotlight, but in recent years interest
has arisen to their application potential, as their extremophilic nature offers unique opportunities in
terms of nanoengineering. Although traditionally recognized for their roles as infectious agents,
viruses have been engineered over the past decade as highly promising nanoplatforms for the
targeted delivery and treatment of human diseases (7). Their attractive features include; naturally
self-assembled nanostructures, biodegradable and the viral capsids provide ideal templates for
engineering multifunctionality, including multivalent display of surface ligands and encapsulation
of therapeutic molecules (8). Plant-based viruses and bacteriophages are typically considered safer
nanoplatforms than mammalian viruses because they cannot proliferate in humans and hence are
less likely to trigger adverse effects (9). Another group of viruses that fits this criterion is archaeal
viruses yet their potential remains untapped. Archaeal viruses offer distinct advantages when
compared to the plant-based viruses and bacteriophages. When designing efficient carrier systems,
7
physical properties such as size, morphology, and surface charge, of the nanocarrier exterior have
direct influence on cellular uptake and intracellular trafficking (7). As a group, archaeal viruses
represent the most diverse morphotypes, providing more sizes and shapes to choose between than
any other virus group. Moreover, lack of stability has been reported to be problematic for synthetic
nanoparticles. The inherent stability of archaeal virus particles provides an array of stable
nanoparticles perfectly suited for harsh industrial processes.
Overall, archaeal viruses appear to be promising nanoplatforms but no single virus has been
assessed either in vitro or in vivo to test it. In order to develop a virus into a therapeutic
nanoplatform in depth knowledge of the virus is essential. First of all, virus characteristics such as
morphology, surface functionalities, replication, and extracellular stability need to be investigated to
provide a basis for future developments. Next, the candidate is studied to determine cellular uptake,
intracellular trafficking and accumulation, together with pharmacokinetics, toxicity and
immunogenicity (7). Together, this knowledge can be used for evaluating if indeed the potential is
worth pursuing. This thesis presents the first in depth investigation of any archaeal virus, SMV1, as
a potential nanoplatform for applications in nanomedicine.
2
Aims of the thesis
This thesis aims at evaluating archaeal viruses as novel nanoplatforms using the two
hyperthermophilic archaeal viruses, SMV1 and SSV2 as model viruses. It further aims at gaining
insights into the viral life-cycle of SMV1 to provide a strong knowledge base for developing SMV1
into a nanovector platform by
− Investigating host range and replication (Chapter I)
− Optimizing purification methods (Chapter I)
− Developing a fluorescent labelling method for SMV1 VNPs (Chapter II)
− Determining extracellular stability of SMV1 (Chapters I and III)
Secondly, it aims at finding potential applications of SMV1 as a novel therapeutic nanovector by
− Investigating the intracellular fate in two endothelial cell lines of human origin (Chapter II)
− Evaluating cell toxicity response (Chapter II)
− Investigating the in vivo stability of the virus particles (Chapter III)
8
− Analysing SMV1 trafficking in the mammalian GI tract (Chapter III)
− Challenging human intestinal organoids to determine the fate of SMV1 in the human gut
epithelium (Chapter III)
3
Archaea: the new branch on the tree
One of the defining events in biology over the last half century was the discovery of the Archaea
and with it a radical reorganization of the tree of life. In the late 1970s, Dr. Carl Woese spearheaded
a study of evolutionary relationships among prokaryotes. He relied on 16S ribosomal RNA
sequences to construct taxonomies that revealed that life, rather than fitting into five kingdoms or a
bipartite eukaryote-prokaryote dichotomy, is best divided into three distinct domains of life (Fig 1)
(1). He discovered that the prokaryotes were actually composed of two very different groups –
the Bacteria and a newly recognized group that he called the Archaea. Each of these groups is as
different from the other as they are from Eukaryotes. Although the archaeal domain was only
formally proposed 25 years ago, they evolved as one of the three primary lineages several billion
years ago. Over this short period of investigative history, the scientific community has discovered
many fascinating things about the Archaea – from evolutionary relationships to biological novelties.
The Archaea are characterized by several distinguishing traits, including ether-linked membrane
lipids, unique cell wall components and the ability of certain genera to produce methane (Fig 1).
But the most striking differences between Archaea and Bacteria are seen in the organization of their
information-processing systems. The structures of ribosomes and chromatin, the presence of
histones, and sequence similarity between proteins involved in translation, transcription, replication
and DNA repair all point to a closer relationship between Archaea and Eukaryotes than between
either of these and Bacteria (10, 11). In contrast many of the metabolic pathways of Archaea more
closely resemble their bacterial rather than eukaryotic counterparts (12, 13). These unique and
shared traits support the status of Archaea as a distinct domain of life with specific connections to
Eukaryotes.
3.1 Extremophilic Archaea
The first characterized Archaea thrived in conditions that were characterized by high salinity, high
temperature and acidity, or strict anoxia – which led to the hypothesis that Archaea require extreme
conditions to thrive. However, the unifying ecological perspective of the Archaea as extremophiles
9
was largely abandoned with the discovery of Archaea throughout the world’s oceans (14). Although
many archaeal species are prevalent in extreme environments and they hold many records for
growth and survival under many ecological extremes, studies have begun to show that they also
compete successfully in “mainstream environments” (15). Further, they contribute significantly to
global nutrient cycling and are much more widespread than previously thought. On the basis of cell
counts and molecular studies Archaea account for as much as 20% of the total biomass in the
oceans (16). Nevertheless, it is the extremophiles that fascinates us and makes catchy headlines with
“living life on the edge”.
Bacteria
Archaea
Eukarya
Ester
Ether
Ester
Metabolism
Bacterial
Bacterial-like
Eukaryotic
Core transcription apparatus
Bacterial
Eukaryotic-like
Eukaryotic
Nucleus
No
No
Yes
Organelles
No
No
Yes
Methanogenesis
No
Yes
No
Pathogens
Yes
No
Yes
Carbon linkage of lipids
Fig. 1⏐Schematic drawing of a phylogenetic tree showing the relative positions of evolutionary
pivotal groups, adapted from Lehninger Principles of Biochemistry, Fifth edition (2008). The table
lists selected traits of the Bacteria, Archaea and Eukarya. The occurrence of a trait is for the
majority of cases (not necessarily absolute).
10
The Archaea are notorious for inhabiting some of the most forbidding places on earth, and thrive
under conditions that few Bacteria and no Eukaryotes would tolerate. BOX 1 provides examples
and descriptions of extreme habitats where Archaea dominate or out-compete Bacteria. In order to
survive under these hostile conditions, archaeal groups have a myriad of genetic and biochemical
adaptations. Rather than having one basic set of adaptations that works for all environments,
Archaea have evolved separate features that are customized for each environment (17). For example
psychrophiles tend to have proteins with a reduced hydrophobic core and a less charged protein
surface to maintain activity under cold temperatures and thermophiles have reverse gyrase, an
enzyme that introduces positive supercoils in DNA and thereby protects it from unwinding (18).
These are just two examples of some of the unique adaptations found in extremophilic Archaea.
Many of the unique features have the potential for applications in biotechnology, of the few
examples in current use, the most well-known is the thermostable enzymes used for DNA
amplification by PCR (Pfu DNA polymerase from Pyrococcus fusriosus). The potential of archaeal
models in biotechnological applications is significant. For example, the ability of methanogenic
Archaea to thrive under anaerobic conditions means that they are ideally suited for use in the
bioremediation of anoxic sludge, such as contaminated shorelines (19). Also, the production of
methane by methanogenic Archaea could be used as a fuel source. Another hot topic is investigating
thermophilic Archaea to find novel thermostable enzymes, which are finding many commercial
uses for example as biocatalysts in the application of ‘green chemistry’ (20). Despite these
promising aspects of Archaea, not much progress or research is focused on these subjects. One
major limitation has been that working with extremophiles is not an easy feat. Organisms that thrive
in boiling acid or anoxic environments are not conducive to routine genetic techniques. And upscaling cultures and workflow is often nearly impossible or simply too expensive. Unfortunately,
this means that numerous biochemical and structural studies on Archaea are not being underpinned
by in vivo data (19). However, progress is being made and in fact many species can be cultivated
with relative ease when their growth conditions are optimized. Consequently, there are now several
model organisms available for extremophilic Archaea; methanogens, halophiles and thermophiles
(21). Among the latter group, there are genetic systems for Sulfolobales and Thermococcales.
Especially, Sulfolobales has been thoroughly investigated and in terms of molecular and
mechanistic details, growth conditions and genetic tools, Sulfolobus is well understood and a good
choice for a ready-to-use model organism (21).
11
BOX 1⏐Examples of extremophilic Archaea and their habitats
Hyperthermophiles
Extremely
thermophilic
(”super
heat-loving”)
Archaea are characterized by growth optima of 80°C
or more, with a current record of growth at 121°C
(22). They colonize volcanic terrestrial environments
and
deep-sea
hydrothermal
vents,
growing
aerobically or anaerobically, and often derive energy
by sulphur reduction.
Extreme halophiles
Halophilic Archaea thrive in environments with salt
concentrations approaching saturation, such as
natural brines, the Dead Sea and hypersaline lakes.
They often have striking pigmentation in red, orange
or purple and they are the possessors of the first
known proton pump, bacteriodhopsin which is driven
just by sunlight (23).
Psychrophiles
Psychrophilic
Archaea
proliferate
at
0-10°C,
metabolize in snow and ice at –20°C and can survive
at –45°C. Most (~75%) of the Earth’s biosphere is
cold, and the largest proportion and greatest diversity
of Archaea exists in cold environments, such as
alpine and polar habitats, the deep ocean, terrestrial
and ocean subsurface, and the upper atmosphere (24)
Methanogens
Methanogens are a group of strictly anaerobic
Archaea that are characterized by the unique ability
to produce methane as a catabolic end product. These
organisms are found in anaerobic marine and
freshwater environments and thrive across a broad
range of temperatures, salinities and pH (2).
12
3.2 Sulfolobus
The genus Sulfolobus belongs to the Crenarchaeota, one of the main archaeal phyla (Fig 2). The
first member, S. acidocaldarius, was isolated in 1972 by T. Brock from a hot spring in Yellowstone
National Park (25). Later on, different members of Sulfolobus were characterized from acidic hot
springs and mud holes all over the world e.g. Iceland, Italy, Japan, New Zealand, Russia and the
USA. The organisms are obligate aerobes that grow optimally between 70-85°C and pH 2-3. In
their natural environment of solfataric hot springs, sulphur compounds are utilized as energy
sources during aerobic respiration, resulting in the production of sulphuric acid. When growing
cultures in the laboratory this results in a characteristic odor. A few Sulfolobus species are reported
to grow chemolithoautotrophically but are easily cultivated aerobiacally under heterotrophic
conditions in the laboratory and exhibit doubling times of 3-6 hours h (21). Interestingly, Sulfolobus
species have been a source for the isolation and characterization of a large number of genetic
elements such as plasmids, conjugative plasmids, and viruses (26, 27). These genetic elements are
being studied intensively and some of them have been used for shuttle vector construction in the
development of genetic tools (28).
Sulfolobus spp. have developed into model organisms for studies of DNA translation, transcription,
and replication, cell division, metabolism, and many other cellular aspects. One reason for this
popularity is that Sulfolobus spp. have been easily adapted to laboratory settings and they can be
cultured to high cell densities (∼1010 cells/mL), see figure 2 for electron microscopy images of a
Fig. 2⏐A1 scanning electron microscopy image. A2 transmission electron microscopy image. Both
images show the irregular coccoid morphology of a typical Sulfolobus cell with the cell envelope
consisting of a lipid membrane and an S-layer (Scale bars, 200 nm) (29).
13
S. islandicus REY15A ΔC1C2
S. solfataricus P2 5E6
Hot spring
Yellowstone National Park
Solfataric fields
Piscarelli, Italy
Growth optima
80°C, pH 2-3, aerobic
80°C, pH 2-3, aerobic
Genome
2.2 Mbp
Circular chromosome
2.9 Mbp
Circular chromosome
CRISPR
Deletion (spontaneous)
Knock-out
Predicted proteins
2819
2977
Associated viruses
SSV2, SIFV, SIRV1, SIRV2,
SMV1
STIV, SSV, SSVK1, SSVRH
Origin
Note-worthy
References
20 whole genomes sequenced of
different S. islandicus strains
(30-33)
More than 10% of the genome
consists of mobile elements
(34-36)
Table 1⏐Characterization of the two archaeal species; S. solfataricus and S. islandicus
typical Sulfolobus cell. Additionally, their proteins are very amenable for obtaining 3D structures,
and the PDB database contains > 500 crystal structures obtained from different Sulfolobus species.
The cell cycle of Sulfolobus is the best-characterized of any archaeal genus and today there are 31
whole genome sequences of four Sulfolobus species available in public databases (NCBI). Together,
this provides a strong basis for using Sulfolobus species to address fundamental questions of
archaeal biology and to use them in biotechnological studies. The present study employ a range of
Sulfolobus species but primarily two strains are used; S. solfataricus P2 5E6 and S. islandicus
REY15A ΔC1C2. Table 1 provides a more in depth description of the two Sulfolobus strains. Both
are well-established laboratory strains, and can be grown on solid as well as in liquid medium. Like
other Sulfolobus cells they do not form chains or aggregates, thus microscopy and flow cytometry
can conveniently be applied (37). Additionally, both strains are hosts for a variety of archaeal
viruses including the viruses investigated in chapters I-III. Lastly, the two strains are both CRISPR
deletion mutants, thus they lack their CRISPR-mediated virus defence (se BOX 2). With their
natural defence deactivated they are highly susceptible to virus infection and are ideal candidates
for virus propagation. All taken together, these two strains represent ideal model organisms for the
present study, investigating and propagating the archaeal viruses, SMV1 and SSV2.
14
BOX 2⏐Features of the CRISPR-Cas adaptive immune system
The CRISPR-Cas system provides immunity against viruses and plasmids, in about 40% of
Bacteria and 90% of Archaea, by targeting the nucleic acids of the invaders. The system is
significantly more complex than the known restriction-modification systems in that the cleavage is
restricted to specific sequences on the invader genome and the system adapts when exposed to new
and returning invading elements. Therefore it has been classified separately from commonly
known defense systems of Bacteria and Archaea as an adaptive immune system.
Overview of the CRISPR-Cas adaptive immune system. The CRISPR-Cas system consists of Cas genes and one or
more CRISPR arrays. The CRISPR-Cas targeting pathway functions in three distinct pathways. Stage 1: CRISPR
spacer acquisition. DNA fragments (protospacers) from an invading plasmid or virus are captured and incorporated
into the CRISPR locus immediately after the leader end on host DNA. A CRISPR array consists of unique spacers
(colored boxes; spacers are numbered sequentially with the most recently acquired spacer having the highest number)
interspaced between repeats (black diamonds). Stage 2: CRISPR expression. CRISPR arrays are transcribed from a
promoter in a leader region and processed into mature crRNAs by Cas proteins. Stage 3: CRISPR interference. crRNA
containing a spacer that has a strong match to incoming foreign nucleic acid (plasmid or virus) initiates a cleavage
event (shown by scissors); Cas proteins are required for this process. DNA cleavage interferes with virus replication or
plasmid activity and imparts immunity to the host. This figure is based on the CRISPR-Cas system in Streptococcus
thermophilus, which represents a well-studied and relatively simple CRISPR-Cas system. From (38).
15
4
The wondrous world of archaeal viruses
Over the past three decades we have gained many insights into the physiology, biochemistry, and
evolutionary relationships of the Archaea. But one of the more fascinating discoveries has been the
wondrous world of archaeal viruses. As a group, archaeal viruses constitute the smallest group of
known viruses. These viruses are often morphologically and genetically unique with exceptional
life-cycle traits (5). To date, all characterized archaeal viruses, infect extremophilic Archaea (Table
1, Table 2, Chapter IV). The majority infects hyperthermophiles belonging to the crenarchaeal
genera Sulfolobus and Acidianus or halophiles of the euryarchaeal genera Haloarcula and
Halorubrum (39). Archaeal viruses have yet to be isolated from non-extreme environments.
However, a putative provirus was recently recognized in the genome of a thaumarchaeon and viruslike particles (VLPs) resembling Archaea-specific viruses have been detected in freshwater
sediments (40, 41). Further, several mesophilic crenarchaeota have been isolated thus the hosts are
there we just need to isolate the viruses. In contrast, bacterial viruses have been isolated from both
extremophiles and mesophiles, and the majority infect the latter. To date, the 6649 isolated bacterial
viruses infect hosts belonging to almost 200 different genera, whereas the 83 archaeal viruses infect
hosts belonging to less than 15 genera. Despite the lack in isolated archaeal viruses, research has
demonstrated that archaeal viruses occupy a distinctive part of the virosphere and display
morphotypes that are not associated with the other two domains of life, Bacteria and Eukarya.
4.1 Morphotypes of archaeal viruses
DNA viruses of the Archaea have highly diverse and often exceptionally complex morphotypes.
Given this diversity, they are often classified into viral families based on their virion shape. So far
16 different morphotypes have been described for archaeal viruses and four of these are unique;
lemon-shape, bottle-shape, droplet-shape, and coil-shape (Fig 3, Fig 4). For bacteriophages, only
nine morphotypes are known and no new morphotypes have been discovered since the 1970s (42).
Although morphologically diverse, all known archaeal genomes are DNA genomes. Moreover,
genome-sequence analyses have demonstrated that most of the archaeal viruses only share sequence
similarity with closely related viruses except for head-tailed isolates (43, 44). The following
sections will present short descriptions of representatives from all 10 archaeal virus families with
special emphasis on the spindle-shaped viruses, as the two archaeal viruses, SMV1 and SSV2, used
16
in the present study belong to this group. The sections are intended to inspire scientists in the
nanotechnology field not yet familiar with the exceptional diversity of archaeal viruses.
4.1.1 Head-tailed virions
Head-tailed viruses (Caudivirales) comprise 96% of known bacteriophages and are by far the most
studied compared to other viral morphotypes (42). All head-tailed archaeal virus isolates are
associated with Euryarchaeota and they exclusively infect extreme halophiles or methanogens that
are either mesophilic or moderately thermophilic. Head-tailed viruses are non-enveloped, and
contain an icosahedral head, which encapsulates the viral genome and is attached to a tail (45). So
far, two thirds of the isolated haloarchaeal viruses represent head-tail morphology whereas only two
head-tailed viruses infecting methanogens have been isolated (Table 1, Table 2, Chapter IV). These
viruses are classified by tail morphology into three families, Myoviridae (long, contractile
tails), Siphoviridae (long, non-contractile tails), and Podoviridae (short tails). The majority of headtailed haloarchaeal viruses are described as lytic viruses e.g. the host cell is lysed to release viral
progeny. However, some head-tailed haloarchaeal viruses are able to exit hosts without causing
lysis (persistent infection) or not cause immediate lysis following entry into the host (temperate
infection) (45). Only three temperate haloarchaeal viruses have been discovered including ΦCh1
(Fig 3), which exists as a chromosomally integrated prophage in Natronobacterium magadii (46).
Although viral diversity has only been studied in a few Archaea-rich habitats, it is becoming
increasingly clear that head-tail particles might be a rare virion morphotype in environments where
Archaea dominate. For example, electron microscopy observations of samples from hypersaline
waters where haloarchaea predominate, spindle-, spherical-, and star-shaped viruses are the most
common observed morphotypes (47). The abundance of head-tailed haloarchaeal viruses among
those isolated clearly reflects a major bias imposed by difficulties in host cultivation and virus
isolation methods favouring lytic viruses that can be observed through the formation of plaques.
4.1.2 Spherical virions
Two types of spherical archaeal viruses have been recognized: (i) those having a helical
nucleoprotein core and (ii) those with an icosahedrally symmetric capsid. The former group is
represented by the viral family Globuloviridae, which includes crenarchaeal Pyrobaculum spherical
virus (PSV), see figure 3 for EM image. The virion consists of a lipid envelope surrounding a linear
17
Fig. 3⏐ Negative contrast electron micrographs of virions of viruses; ABV, APBV1, SNDV, AFV1,
SIRV2, PSV, ΦCh1, and ACV. Scale bars = 100 nm. Images adapted from (46, 48-52).
18
double-stranded DNA (dsDNA) genome, which is tightly packed into a helical nucleoprotein core.
The second group of spherical archaeal viruses comprises four halophilic and two
hyperthermophilic viruses. Two new viral families have been proposed separating the
hyperthermophilic viruses (STIV and STIV2) into the “Turriviridae” and the halophilic viruses
(SH1, HHIV2, PH1, and SNJ1) into the “Sphaerolipoviridae”. The icosahedral virions are nonenveloped with an inner lipid layer.
Despite their shared morphotype the spherical viral families do not necessarily share more features
with each other than with other archaeal virus families. For example, SH1 is a lytic virus with a
high burst-size of approximately 200 virus particles per cell, and the release mechanism is based on
cell disruption (53). By contrast, STIV is non-lytic and persists stably in the host cell. In line with
differences in morphology and virus-host interactions, the genomes of the two viruses reveal no
evidence of homologous gene content. Furthermore, although the genome of STIV is circular, that
of SH1 is linear (5).
4.1.3 Linear virions
Linear viruses represent the most abundant virion morphotype in extreme geothermal environments
and have also been observed in hypersaline waters (5, 54). All species isolated from these
environments infect Crenarchaeota and carry dsDNA genomes, a property not previously observed
for any linear virus. These viruses can be divided into four groups: (i) filamentous, (ii) rod-shaped,
(iii) coil-shaped, and (iv) bacilliform (Table 1, Chapter IV). Originally, the discrimination of the
families was based mainly on differences in virion structure but later it was supported by
comparative genomics (5). Filamentous and rod-shaped viruses are classified into the family
Lipothrixviridae and Rudiviridae, respectively. Lipothriviruses typically have flexible, filamentous
virions that are surrounded by envelopes containing lipids that have been acquired from the host.
They show considerable diversity in their terminal structures, an example is AFV1 (Fig 3), which
carry exceptional claw-like structures (51). Rudiviruses have non-enveloped rigid virions that vary
considerably in length, with the length proportional to the size of the linear dsDNA. To date, five
rudiviruses have been sequenced, including SIRV2 (Fig 3). SIRV2 is a lytic virus that is released by
a unique release mechanism involving the generation of pyramidal structures that, by opening out,
cause a local disruption of the cell envelope and allow virion escape (29).
19
The latter two groups have only one member each. Virions of Aeropyrum pernix bacilliform virus 1
(APBV1, Fig 3) are stiff and bacilli-like, and the virus has been classified into a new viral family,
Clavaviridae. Interestingly, APBV1 infection causes neither retardation of host growth nor lysis or
host cells, and integration of the viral genome into the host chromosome has not been detected (49).
Aeropyrum coil-shaped virus (ACV, Fig 3), which is classified into the proposed family
Spiraviridae, has hollow cylinder-like virions formed by a coiling nucleoprotein fiber. The genome
of APBV1 (5.2 kp) is the smallest known prokaryotic dsDNA viral genome, whereas ACV has the
largest known single-stranded DNA (ssDNA) genome (24.9 kp) (49, 55).
4.1.4 Pleomorphic virions
Recently, a comparative biological and structural study of pleomorphic viruses led to the proposal
of the viral family, ”Pleolipoviridae”. The pleolipoviruses all infect the haloarchaeal Halorubrum
sp. and are defined by similar life-cycles, protein and lipid patterns, and virion organizations (56).
Pleolipoviruses are more or less spherical and distinctively, all the viral particles show spikes
decorating the surface of the viral membrane. The spikes appear to have a random distribution and
are thought to play a role in host interaction (56). The genome of pleolipoviruses resides inside a
lipid envelope without forming any ordered nucleoprotein complex. These viruses are non-lytic,
only retarding host growth, and they acquire their lipids unselectively from the host cell membrane.
Consequently, it has been proposed that pleolipoviruses use budding to exit infected cells (57, 58).
4.1.5 Bottle-shaped and droplet-shaped virions
The virions of the archaeal viruses, Acidianus bottle-shaped virus (ABV) and Sulfolobus
neozealandicus droplet-shaped virus (SNDV), have exceptional morphological features that are so
unique that each of the viruses have been assigned to a new family, Ampullaviridae and
Guttaviridae, respectively. The virion of the ampullavirus ABV is structurally one of the most
complex virions in the viral world and the virions have no elements of icosahedral or helical
symmetry (Fig 3). At the broader end of the bottle-shaped particle are 20 short rigid filaments, their
function remains unclear. The virion of the guttavirus SNDV exhibits a beehive-like capsid with a
pointed end densely covered with thin fibres (Fig 3). The virion surface seems to be either helically
ribbed or a stack of rings (59). This virus presently does not exist in culture collections.
20
4.1.6 Viruses with spindle-shaped virions
Viruses with spindle-shaped virions, single or two-tailed, are common in, and exclusive to the
Archaea (5). They constitute a large fraction of the known archaeal viruses and have been observed
in diverse Archaea-rich habitats, including terrestrial and marine extreme thermal environments and
hypersaline waters (47, 60). Nineteen spindle-shaped viruses have been isolated (Table 1, Table 2,
Chapter IV) and all of them, except for one, infect hyperthermophiles. Spindle-shaped viruses are
diverse in their structural and genomic characteristics, and they have been classified into the
Fuselloviridae family, the proposed “Bicaudaviridae” and “Monocaudaviridae” families and the
genus Salterprovirus, four remain unclassified. Most have a circular genome and carry an integrase
gene that can facilitate integration into host chromosomes (61).
The Salterprovirus, His1, is the only spindle-shaped haloarchaeovirus isolated to date, despite this
morphotype being the most abundant in several hypersaline environments (47). It infects an
extremely halophilic euryarchaeon, Haloarcula hispanica, and morphologically resembles
fuselloviruses. However, unlike fuselloviruses, which have a circular dsDNA genome, His1 has a
linear dsDNA genome. Studies have shown His1 virions to be stable under a range of biochemical
treatments, illustrating the extremely stable nature of His1 virions (62). Acidianus two-tailed virus
(ATV), the only member in the family Bicaudaviridae, has an extraordinary capability to undergo
major morphological changes that occur outside of and independent from its host (63). ATV virions
are released from host cells as lemon-shaped tail-less particles, and when incubated at temperatures
close to the natural infection conditions, long tails develop at each pointed end (52).
In the present study the only member of Monocauaviridae, SMV1, is used as a model organism.
SMV1 was selected based on its unique morphotype, ease of production, and it can be enumerated
by plaque assay. The spindle-shaped virions show considerable size variation and in virus
preparations virions with no tail, one-tail, and two-tails are observed, showing varying degrees of
tail-development (Fig 4). The specific mechanism for tail-development is unknown, however both
temperatures >75°C and proteinase K can trigger extracellular tail-development. SMV1 has a
genome size of 48.8 kb with 51 putative ORFs (64).
21
Fig. 4⏐ Negative contrast electron micrographs of SMV1 and SSV2 virions. SMV1 particles
isolated immediately after release occur tailless: a. SMV1 develops 1 or 2 tails outside of the host
cell: b, c. Only one pole appears to have short tail fibres which can attach to other tail fibres to form
characteristic rosettes: b. SSV2 particles are very uniform and the virus preparations often show a
high degree of virus aggregation into rosette-like structures: d, e. For comparison, an image of a
mixed sample of SMV1 (the large particle) and SSV2 (smaller particles) is shown: f.
22
Twelve known species of the fuselloviruses (Table 1, Chapter IV) can propagate in both major
culturable genera from aerobic, acidic hot springs, Sulfolobus and Acidianus. The virions, which are
in the size range 55–60 × 80–100 nm, have short tails which are uniform in size and carry thin
terminal fibres, which appear to be extremely sticky and readily attach to cellular fragments (65,
66). In addition, they can attach to the same fibres in other virions, leading to the formation of
rosette-like aggregates. The best-studied is the type species of the family, the Sulfolobus spindleshaped virus 1 (SSV1). Its circular genome is positively supercoiled and during infection it
integrates into a tRNA gene in the host chromosome. SSV1 replication can be induced by UV
irradiation or mytomycin treatment (67). Several more SSV viruses have been isolated from
different locations in the world, including SSV2, which were originally isolated from a solfataric
hot spring in Reykjanes, Iceland (31). SSV2 is the second model virus used in the present study and
was selected to complement SMV1, as a similar but smaller morphotype. For comparison of the two
model viruses see table 2 and figure 4. SSV2 bears significant resemblance to SSV1 both in shape
and in genome organization. SSV2 has a genome size of 14.7 kb with 34 putative ORFs (31, 68).
However, initiation of viral replication appears to be regulated differently in the two viruses. While
SSV1 DNA replication in the infected host cell is strongly induced in response to UV irradiation
SSV2 replication in the natural host is induced concurrently with the halt of host growth in a late
growth phase, making it easy to obtain high yields (69). Both SSV2 and SMV1 are well-established
laboratory strains at the Danish Archaea, together with their unique morphotypes, it made them
ideal candidates for the exploration into applications of archaeal viruses.
Monocaudavirus SMV1
Fusellovirus SSV2
Yellowstone National Park
Iceland
Isolation host
Sulfolobus sp.
Sulfolobus solfataricus
Growth optima
75-80°C, pH 3
80°C, pH 3
Morphotype
Spindle-shape
Spindle-shape
1-2 tails, tail fibers at one end
fibers at one pole
Virion diameter
100-120nm
60-90nm
Genome / ORFs
48.8 kb / 51
14.7 kb / 34
Extremely stable virions
Very uniform virions
(64)
(31, 68)
Origin
Note-worthy
References
Table 2⏐Comparison of the two hyperthermophilic archaeal viruses; SMV1 and SSV2.
23
4.2 Application potential of archaeal viruses
Despite their lack in isolated numbers, archaeal viruses have revealed many surprises and changed
our understanding of the virosphere. These include the extraordinary virion morphotypes described
above, unique life-cycle characteristics, and a better understanding of the evolutionary relationship
between viruses. Initially, the unique morphotypes and evolutionary significance stole the spotlight,
but in recent years interest has arisen to the application potential of archaeal viruses (5). All isolated
archaeal viruses are extremophilic in nature and show adaptations to the extreme environments of
their host, making them extremely stable nanoparticles (particle size in the range of 1-100nm).
Thus, viruses of extreme halophiles show stability in solutions of high salt concentration (∼3–5M),
similarly, viruses infecting acidophilic hyperthermophiles are stable under very aggressive
conditions, at pH values lower than 3 and temperatures above 80°C. These stability features offer
attractive traits for exploration of their potential in bionanotechnological applications (70, 71).
Although traditionally recognized for their roles as infectious agents, viruses show commercial
value in bionanotechnological applications. For example, viruses have been engineered over the
past decade as highly promising nanoplatforms for the targeted delivery and treatment of human
diseases (7). A broad range of genetic and chemical engineering methods have been established that
allow viral particles to carry large payloads of imaging reagents or drugs to specific target sites (7).
In general, plant viruses and bacteriophages are considered safer in humans than mammalian
viruses (even replication-deficient strains of mammalian viruses are potentially pathogenic and/or
cytotoxic) (72). Viral nanoplatforms based on plant viruses or bacteriophages are less likely to
trigger negative downstream effects in mammals due to an inability to proliferate, also preclinical
studies have shown that plant viruses can be administered at doses of up to 100 mg (1016 VNPs) per
kilogram bodyweight without clinical toxicity (73). In contrast, 1011 Adenovirus particles can cause
severe hepatoxicity (74). Archaeal viruses should be included in the “safe” virus group of
prospective viral nanoplatforms, as they cannot proliferate in mammalian cells either. Moreover, no
pathogen of archaeal origin has been identified (75). However, only few studies have looked into
the application potential of archaeal viruses, despite their promise of superior stability.
24
1
2
4.2.1 SIRV2 and His1
SIRV2 is a hyperthermophilic rod-shaped archaeal virus (Fig 3) that has been studied as a potential
nanobuilding block by Steintmetz et al. (2008). SIRV2 was shown to be an exceptionally robust
nanoparticle, stable at extremes of temperature and acidity as well as in diverse non-natural harsh
conditions (70). Further, their studies showed that, depending on the chemistry and hence
attachment site used, modifications in form of biotin labels could be targeted specifically to the
virion body and at the ends (Fig 5). The fact that native SIRV2 offers three different chemical
attachment sites is unique for rod-shaped viruses and opens up possibilities for differential labelling
with different physical properties. Thus, SIRV2 could be engineered into a multifunctional scaffold
for future applications in nanoscale templating, nanoelectronics and biomedical applications (70).
Another study has tested the inherent stability of the halophilic virus His1. His1 virions have
different sizes, but prove to be extremely stable under various biochemical treatments, such as
detergents, solvents, proteases etc. providing one more example of the inherent stability of archaeal
virus particles (62). These studies are small first steps in evaluating the application potential of
archaeal viruses, much more work is needed before a “useful” archaeal virus is developed.
5
When viruses become useful
Twenty years ago most people (including many scientists) thought of Bacteria solely as agents of
disease, best treated with soaps, detergents and antibiotics (76). Today, most of us are aware that
Bacteria play crucial roles not only in the environment but also in our own bodies. We now
understand that microorganisms make up almost 90% of the cells in our bodies, and play a critical
A
B
Fig. 5⏐ Negative contrast electron micrographs of SIRV2. Biotinylated SIRV2 particles labelled
using carboxylate-selective chemistry, labelling to both the virion body and ends: a. Biotinylated
SIRV2 particles labelled using amine-selective bioconjugation techniques, labelling occurred
exclusively at the virion ends: b. Adapted from (70).
25
role in digestion and the immune response (77). In plants, microorganisms form important
mutualistic relationships, providing nitrogen fixation and defence against pathogens and much more
(78). We are beginning to realize that we need our good microbes and there is an increased focus on
identifying them. However viruses are generally not included in the beneficial microbe list. But
recent work indicates that animal viruses in the gut are not just pathogens that infect host cells and
cause gastroenteritis but also are symbiotic modulators of host physiology. Thus giving rise to the
question of the existence of a ”good” gut virome. Further, a study by Venkataraman et al. (2008)
revealed that ‘Seneca Valley Virus-001’ demonstrates cancer-killing specificity that is 10.000 times
higher than that seen in traditional chemotherapeutics, with no overt toxicity (79). Thereby, it is
reasonable to speculate that beneficial viruses exits, all we need to do is find them (80).
5.1 Commercial development
Another definition of beneficial viruses is a more commercial viewpoint, where beneficial viruses
are seen as viruses that provide a trait that has industrial value. Some of the best-characterized
viruses of commercial value in plants, are those that enhance the beauty of ornamental plants, such
as Tulip breaking virus that cause flame-like streaks on the petals (76). Another important use of
viruses, both medically and commercially, is in vaccine development. Vaccination is the most
effective medical intervention against diseases caused by human viral pathogens. There are two
basic types of virus vaccines: live attenuated and inactivated. Live attenuated vaccines are produced
by modifying ”live” viruses into a weakened/altered state that retains the ability to replicate and
produce immunity, but usually not cause disease. Inactivated vaccines are produced by killing the
disease-causing viruses with chemicals, heat, or radiation (81). They are considered safer than live
vaccines as they cannot mutate back to their disease-causing state. Several more vaccine strategies
exists including, subunit-, toxoid-, conjugate-, and DNA-vaccines. In the fight against infections,
viruses can also be used to kill harmful Bacteria in plants and humans. With more Bacteria
becoming resistant to the most commonly used antibiotics, the use of viruses as antibacterial agents
could prove a valuable alternative (82). Last, viruses are being used as nanoplatforms with
applications in materials science and medicine. The main advantages of utilizing viral nanoparticles
(VNPs) are their nanometer-range size, their propensity to self-assemble, their high degree of
symmetry and polyvalency, the relative ease of producing large quantities of material, their
exceptional stability, robustness, and biocompatibility (70). The development of viruses and
bacteriophages into nanoplatforms is a highly promising field with new viruses being explored as
26
potential nanoplatforms each year. The next section will cover some of the major design strategies
when developing novel viral nanoplatforms.
6
Design strategies of viral nanoplatforms
The ability to control and target drug delivery is crucial for effective and efficient medical
treatment; however, the lack of control over site-specific localization and, hence, bioavailability at
the desired site still remains one of the major challenges in modern-day medicine (7). Over the last
decade, viral nanotechnology have tried to address this problem by developing viral nanocarriers
targeting specific locations in the human body. The ability to form a well-defined closed structure
that can subsequently disassemble offers a convenient and powerful strategy for the controlled
encapsulation and release of various functional therapeutics, such as proteins, enzymes, and
polymers (9). As the exteriors of VNPs are composed entirely of proteins, by functionalizing their
surfaces with appropriate ligands, viral nanocarriers can be targeted to specific cells and locations
within the body. With regard to the design of efficient carrier systems, the physical properties, such
as size, shape, surface chemistry and composition, of the nanocarrier exterior have direct effects on
cellular uptake, intracellular distribution and accumulation, retention and excretion times (83). As
shown in figure 6, these numerous variables lead to a large combination of potential nanoparticles
that could be developed; therefore a semi-rational approach is required when developing viral
nanoplatforms.
The impact of particle size on nanocarrier function has been well studied. Size influences almost
every aspect of particle function including degradation, flow properties, clearance, uptake
mechanism, and biodistribution (84). The diameter of particles administered in blood vessels,
airways or gastrointestinal (GI) tract dictates their velocity, diffusion and adhesion to walls.
Movement of particles in tissues, whether arriving by migration or injection, is also limited by size
due to steric hindrance in the extracellular matrix. Another important factor is surface chemistry,
which primarily influences the interactions of particles with cells and tissues in the body. For
example, it has been shown that strong positive or negative surface charges lead to rapid clearance
of nanoparticles, whereas a slight negative charge lead to a lesser amount of phagocytosis (85). In
addition to size and surface characteristics, the shape of nanoparticles has been identified as a key
factor in carrier performance. Traditionally, nanoparticles used as nanocarriers have been spherical
27
Fig. 6⏐ Overview of the main physical properties that play important roles in the cellular uptake,
intracellular distributions and accumulation, retention and excretion times of nanocarriers. From (7).
in shape. However, several non-spherical nanoparticles such as gold nanorods and carbon nanotubes
as well as other structures with high aspect ratios are now being investigated. In short, several
studies show that nanoparticles with cylindrical-, rod-, lamella-, and disk-shapes display higher
circulation times than spherical equivalents. Further, elongated nanoparticles show higher efficiency
in adhering to cells compared to spherical nanoparticles. The curve shape of spherical nanoparticles
allows a limited number of their binding sites to interact with target cell receptors. While, the
elongated nanoparticles have a higher surface area that facilities the multivalent interaction of these
particles with the cell surface. As a result, the elongated nanoparticles with higher aspect ratios
show higher uptake than their spherical counterparts. To fabricate nanoparticles into complex
shapes can be technically challenging. Thus, scientists have turned to viruses to obtain distinctive
non-spherical shapes for the design of nanoplatforms. Here, archaeal viruses can provide unique
shapes not found among other virus groups (Fig 3).
28
6.2 Production
Viruses demonstrate a remarkable plasticity in their metastable structure and dynamics, including
coordinated assembly and disassembly (8). This propensity to self-assemble with atomic precision
is a major advantage, as no man-made steps need to be applied in order to assemble the VNPs.
When choosing a virus as a potential nanoplatform, the first issue to be addressed is that of
production. An ideal viral nanoplatform should be one that can be produced in a highly
concentrated form, using a convenient and reproducible production scheme (86). The nature of
virus biology will usually determine the means of production, thus basic knowledge of the viral lifecycle is essential when choosing a potential candidate. Whether a virus has a lysogenic or lytic lifecycle is quite important. A lytic virus will lyse the host cells to liberate the viral particles and
accumulate in the culture medium. Here, it is important to know at which time after infection the
new viruses burst from the host cells, as to get high yields. With lysogenic viruses, viral replication
needs to be induced. Some lysogenic viruses have been shown to be induced by UV irradiation or
mytomycin treatment (67, 87). However, the optimal induction conditions differ from virus to virus,
thus finding the best method is key. Another production method uses heterologous expression
systems such as E. coli or yeast to produce non-natural VNPs. Typically, viral structural protein
genes are cloned into a plasmid vector under strong promotors, allowing for a high level of
production of the recombinant proteins, which in many cases have been shown to self-assemble
despite the lack of nucleic acids, forming empty capsids (88, 89). Consequently, the distinction
between viral nanoparticles (VNPs) and viral like particles (VLPs) is made.
6.3 Genetic engineering
VNPs and VLPs are assembled into the virion body from protein subunits whose structure and
physicochemical properties can be modified by genetic engineering. For example, viruses can be
engineered to exhibit functionalities through the insertion of bioactive peptides and proteins into the
capsid. The peptides and proteins can be completely foreign to the viruses and can serve as handles
for bioconjugation, introduce peptide-based affinity tags and insert peptides as targeting ligands or
as epitopes to stimulate the immune response (vaccine development) (9). Figure 7 illustrates the
scheme for site-specific modification of Adeno-associated viruses via a genetically engineered
aldehyde-tag (90). Viable genetic modification strategies require a capsid site that fulfils the
following criteria: i) tolerance to the insertion, not disrupting important virus properties, such as
29
Fig. 7⏐ Adeno-associated virus 2 (AAV2) can be site-specifically modified by genetically encoded
aldehyde tags. A 13-amino-acid sequence can be inserted into the cap gene without loss of virus
function and infectivity. The inserted sequence is recognized by cellular Formylglycine generating
enzyme, which converts cysteine to an aldehyde-bearing formylglycine residue: a. The aldehyde tag
can then be covalently conjugated with hydrazide- or hydroxylamine-functionalized
targeting/labelling elements, such as peptides or antibodies: b. Adapted from (90)
capsid assembly, genome packaging and infectivity ii) maintenance of the desired functionality of
the inserted domain iii) if appropriate, suppression of any undesired innate virus properties (e.g.
natural tropism) (91). Such ideal capsid insertion sites can be identified through comparative
structural studies and systematic mutagenesis efforts (92, 93).
In general, prokaryotic viruses have very low uptake efficiency by mammalian cells, but can be
adapted to bind mammalian receptors by genetically engineering a eukaryotic ligand into their
capsid. A study by Hajitou et al. (2006) reported the construction of a hybrid vector that comprises
an Adenovirus cassette inserted into the genome of a single-stranded bacteriophage, resulting in a
peptide motif that is displayed on the phage capsid. The peptide binds to αv integrins, a cell-surface
receptor overexpressed in both tumor and endothelial cells. Thus, the hybrid has potential as a
nanoplatform for ligand-directed tumor targeting (94).
30
6.4 Chemical engineering
VNP and VLP capsid proteins can also be chemically modified using bioconjugation protocols.
Targeting of amino acids with reactive side chains such as thiols, amines, carboxylates or other
functional groups in viral capsids, either naturally occurring or introduced in them by genetic
engineering, has allowed the chemical coupling of small molecules to the capsid. Modification
strategies include functionalization with antibodies, polymers, peptides, biotin, fluorescent reagents
and drugs using N-hydroxysuccinimidyl ester (NHS), maleimide, isothiocyanate and carbodiimide
chemistries (9). In addition, these methods can be used to incorporate synthetic functional groups to
allow the use of click chemistry. Click reactions are favoured because they proceed rapidly with
high selectivity, specificity, and yield. Moreover, click reactions are bio-orthogonal; neither the
reactants nor their product's functional groups interact with functionalized biomolecules. Second,
the reactions proceed with ease under mild non-toxic conditions, such as at room temperature and,
usually, in water (95). The most popular click reaction is the azide-alkyne cycloaddition, which
fuses together two unsaturated reactants (azides and alkynes). This click reaction is unique in that
the azide moiety is absent in almost all naturally existing compounds, lacks reactivity with natural
biomolecules, and, consequently, only undergoes ligation with a limited set of partners such as
activated alkynes. The copper-catalyzed azide-alkyne cycloaddition (CuAAC) protocol has been
used for the attachment of a wide variety of ligands to the surfaces of VNPs (95). Figure 8
represents a bioconjugation scheme using both NHS and CuAAC techniques for the modification of
VNPs and VLPs.
Fig. 8⏐ The functional groups (primary amines) of lysines, exposed at the surface of the viral
capsid, are converted to alkynes by a reaction with NHS ester. The alkyne modified capsid can react
with azide derivatized molecules for further functionalization via copper-catalyzed azide-alkyne
cycloaddition (CuACC) reactions. This scheme is used for the direct coupling of small molecules
such as fluorescent probes to the surface of VNPs and VLPs (96).
31
Significant attention has been paid to chemical modifications of the particle surface so as to
minimize recognition by components of the immune system [10]. PEGylation (the decoration of
PEG onto the surface of nanoparticles) is a common approach to reduce protein absorption, improve
stability and prolong in vivo circulation time (97). A study demonstrated that PEGylation of cowpea
mosaic virus (CPMV) prevents the virus’ interaction with endothelial cells. The PEG molecules
were conjugated to the surface-exposed lysine residues on the CPMV capsid through NHS ester
coupling. It was estimated only ∼22 molecules of PEG1000 and ∼27 molecules of PEG2000 need
to be attached per VNP to eliminate undesired cellular interactions (98).
7
From engineering to applications in medicine
The unique properties of carefully designed viral nanovectors hold great potential for the treatment
of human diseases. The ultimate goal of nanomedical engineering is to develop nanoparticles that
migrate to site-specific locations in the body and exert their therapeutic effect there without causing
any adverse effects. Based on the choice of genetic and chemical engineering a specific
functionalization can be expected. However, One of the major challenges and limitations is that
until viral nanovectors are tested in vivo, their immunological effects cannot be predicted in
advance (99). On the road toward clinical translation of any VNP platform, a detailed understanding
of the body’s response is required. This includes understanding the pharmacokinetics, blood
compatibility, biodistribution, toxicity and clearance properties of the VNPs. Several studies have
shown the functionalization of VNPs but the fate of most of those particles in vivo has not been
investigated, hence there is still a significant lack of knowledge that currently prevents their rapid
implementation as nanomedicine (7).
Toxicity has certainly been a challenge when dealing with viral human pathogens such as Adenoassociated virus. Even when using replication-deficient strains, an inflammatory response is
induced (100). Unlike eukaryotic viruses, viruses derived from bacteriophages and plants are
generally considered to be safe for human treatment. Animal studies have been carried out for a
range of plant-based viruses such as CPMV and Cowpea chlorotic mottle virus (CCMV), as well as
for bacteriophages Qβ and M13, and despite a broad biodistribution, no toxicity was observed (99,
101). Besides toxicity and biodistribution, it is also important to determine the pharmacokinetic
characteristics of the VLPs / VNPs. There needs to be an appropriate balance between tissue
penetration/accumulation and systemic clearance. Longer circulation times allow drugs and
32
reagents to accumulate in target tissues, but the risk of toxicity and stability issues are higher (102).
Pharmacokinetic properties are dependent on the composition of the VNPs such as surface charge
and surface modifications such as PEGylation (see above).
Finally, it should be mentioned that in the recent Human Microbiome Project it has become
apparent that certain Archaea, Bacteria and viruses live symbiotically in the human gut and skin and
play an essential role in maintaining our health. Therefore, when designing viruses as nanocarriers,
it is essential to consider the possible impact on the human microbiome (7).
8
Summary of results
This project aims at evaluating archaeal viruses as novel nanoplatforms with the hyperthermophilic
viruses, SMV1 and SSV2 as models. When developing viruses for applications in nanomedicine it
is essential to gain a comprehensive understanding of their properties from replication to in vivo
behaviour, and particularly any potential toxic effects (99). An overview of the general assembly
line when developing viral nanoplatforms are presented in figure 9 using SMV1 as an example. It
was outside the scope of this project to pursue all three focus areas presented in figure 7. Thus,
throughout this project there have been two major areas of investigation. One area has comprised of
classical virology studies such as life-cycle experiments, host identification and optimization of
purification methods; all to expand our knowledge of the hyperthermophilic archaeal virus SMV1.
This provided a strong foundation for the second area of investigation; studying the interaction
between SMV1 and eukaryotic cells in vitro and in vivo. Importantly, providing us with results to
evaluate its potential as a novel therapeutic nanoplatform. All the experimental work done in the
course of this project, led to three research papers, the major results are presented below.
8.1 SMV1, an ideal model virus of the spindle-shaped kind
When I started my PhD project, SMV1 was a newly isolated virus and it proved to be an easy virus
to work with in the lab e.g. high yields, very stable and could be enumerated by plaque assay, all
very desirable features when selecting a virus for experiments needing large quantities of virus
particles. However, due to it being a “new” virus, it was not well characterized. Therefore, an
important first step was to characterize SMV1. My TEM observations suggests that SMV1 virions
enter into host cells via a fusion entry mechanism, involving three distinct stages; attachment,
33
alignment, and fusion. Upon infection, the intracellular replication cycle lasts 8 h at which point the
virus particles are released as spindle-shaped tailless particles. Replication of the virus retarded host
growth but did not cause lysis of the host cells. As with ATV, SMV1 shows remarkable virion
plasticity by developing a tail/tails independent of the host. An extraordinary feature only observed
Fig. 9⏐ Viral nanomedicine – the assembly line. 1) VNPs can be produced by their natural hosts or
in heterologous expression systems. 2) Once purified, chemical tuning and design is carried out to
confer functionalities. 3) The Functionalized VNPs can then be evaluated in vitro and in vivo.
34
for these two hyperthermophilic spindle-shaped viruses. Additional viral traits are discussed in
Chapter I. In general, SMV1 particles demonstrate an exceptional resilience to a range of harsh
conditions, especially at extremes of temperature and UV irradiance. We hypothesize that the
rosette-like aggregates observed for SMV1 and other spindle-shaped viruses act as a natural defence
mechanism, thus increasing their resilience when exposed to harsh environmental conditions. The
inherent robustness of SMV1 nanoparticles is a very desirable trait in terms of nanoengineering and
emphasizes the importance of evaluating archaeal viruses as potential nanoplatforms for
applications in material science and nanomedicine.
The next steps towards evaluating SMV1 as a potential nanoplatform demanded highly purified
virus suspensions. Traditionally CsCl and sucrose gradients have been used to purify viruses.
However, both methods require long ultracentrifugation time and include a dialyze step before
infectivity can be measured. Also, CsCl can exert toxic effects in downstream cell experiments if
not completely removed and sucrose can also be problematic due to high viscosity and interaction
with viral glycoproteins. Therefore, an alternative method was pursued. Iodixanol (OptiPrep™)
gradients have been used successfully in the purification of various viruses such as HIV, AAV and
hepatitis C virus and it has desirable traits; non-ionic and non-toxic to cells and metabolically inert
(103). I found it to be a superior gradient solution in terms of yield, virus recovery and time. As a
result, SSV2 particles were purified using the same method.
8.2 Recognition of archaeal viruses by human cells
To date, no other study has investigated the interaction between any archaeal virus and cells of
human origin. Therefore, it was important at an early point of the project to investigate the uptake
and intracellular fate of SMV1 and SSV2 in vitro. As these results would guide the design of
following experiments. For example it was important to establish if the VNPs would be taken up or
they would remain in the medium surrounding the cells; each scenario having different
experimental approaches. In order to study the trafficking of VNPs, a fluorescent labeling technique
suitable for SMV1 and SSV2 was required. The green fluorescent general membrane dye PKH67
proved to be an easy labeling technique suitable for both viruses. Next, the fluorescently labeled
SMV1 and SSV2 particles were studied in two different endothelial cell lines of human origin,
hCMEC/D3 and HUVEC. In Chapter II, live-cell microscopy demonstrated viral internalization in
both cell types. SMV1 favorably interacted with both hCMEC/D3 and HUVEC cells in a
35
concentration-dependent manner whereas SSV2 poorly recognizes hCMEC/D3 cells even at high
concentrations. These are the first results demonstrating that archaeal viruses can be taken up and
internalized by human cells, thus indicating a potential as delivery agents.
When developing viruses for applications in biomedicine it is essential to investigate any potential
toxic effects. Neither SMV1, nor SSV2 induced any detrimental effect on plasma membrane or
mitochondrial oxidative phosphorylation. However, quantification of complement activation
products SC5b-9 (Fig 10A) and C5a (Fig 10B) were highly elevated for both SMV1 and SSV2 even
compared to Zymosan, the gold standard to induce complement activation in vitro. The complement
system (CS) is responsible for recognition, elimination and destruction of pathogens and the
activation of the CS triggers the release of proinflammatory molecules such as SC5b-9 and C5a.
Many VNPs have been shown to trigger CS activation, simply due to the fact that they are “foreign”
to the body (104). Briefly, activation of the CS can be harmful if the VNPs enter the systemic
circulation because this may lead to hypersensitivity reactions and anaphylaxis (105). The fact that
both SMV1 and SSV2 are potent triggers of the CS are “bad news” from a nanomedical
perspective. Either the VNPs need to be functionalized to avoid CS activation or a different
approach is needed, one where they avoid systemic circulation.
8.3 SMV1 is extremely stable in the mammalian GI tract
The initial idea of using SMV1 and SSV2 as nanocarriers to target specific locations in the body by
systemic delivery (blood circulation) was largely abandoned by the CS activation results. However,
the inherent stability observed for SMV1 particles prompted a new line of research investigating
SMV1 particles as potential nanocarriers targeting the gut microbiome. The importance of the gut
microbiome for human health has been established and microbiome alterations have been linked to
several diseases. Therefore, the gut microbiome is a potential new target for drug therapy. In order
for orally administered drugs to reach the gut microbiome, they must be stable in the extremely
acidic and proteolytic environment of the GI tract. To overcome this challenge, we investigated
SMV1 as a candidate therapeutic nanovector for the distal mammalian GI tract microbiome.
Stability experiments in Chapter III showed SMV1 to be highly stable in both simulated
conditions of the human GI tract (in vitro) and when passaged orally in mice (in vivo). Importantly,
SMV1 virus could not be detected in tissues outside the murine GI tract indicating that the VNPs do
not enter systemic circulation upon oral administration. In general, high doses of SMV1 elicited a
36
nearly undetectable murine inflammatory response and challenged mice showed no observable
signs of pain or distress.
Other viral vectors have been developed for therapeutic delivery to the distal GI tract and we
compared SMV1 stability to the traditionally used Inovirus, M13KE. SMV1 outperformed this
state-of-the-art vector as measured by in vitro and in vivo survival (replication). In addition, we
found that SMV1 was just as immunotolerant in mice as M13KE. Our results provide strong
evidence that SMV1 in particular and archaeal viruses in general have intrinsically favorable in vivo
characteristics for bioengineering applications, such as drug delivery in the GI tract.
Fig. 10⏐ Concentration dependent complement activation in typical human serum from a healthy
Caucasian individual after incubation with SMV1 and SSV2. Zymosan (200 µg/mL) was used as a
positive control for monitoring complement activation and the blank is the virus storage buffer
(10mM Tris AC, pH6). Complement activation marker (A) SC5b-9; (B) C5a.
37
9
Outlook
The work presented in the following chapters represents the first tentative steps towards developing
archaeal viruses for applications in nanomedicine. Most studies investigating functionalized VNPs
have focused on the in vitro behaviour with only limited data on the performance of engineered
VNPs in vivo. A major challenge and limitation in the development of viruses as nanomedicine is
the lack of in vivo data (7). Until viral nanoplatforms are tested in vivo, their immunological effects
cannot be predicted and hence there is a significant lack of knowledge that currently prevents their
implementation as nanomedicines. Therefore, it was important to include in vivo experiments as
part of this study. However, the in vivo data is based on a very small group of mice and only wildtype and fluorescently labelled SMV1 particles were investigated. Any functionalization of SMV1
particles as part of a therapeutic design strategy would have to include additional in vivo
experiments to ensure no immunological effects of the modified VNPs.
The goal of functionalization is to develop VNPs that migrate to where they are intended to go and
exert therapeutic effect there. The only functionalization included in this study was the fluorescent
labelling of SMV1 and SSV2 VNPs. Thus future application ideas would have to chemically and/or
genetically modify the VNPs for their intended use. The capsid protein structure is not known for
either virus. However, bioinformatic prediction of the 3D-structure shows the presence of several
surface functional groups on the viral capsids available for chemoselective modification. And
although no attempts have been made to genetically modify SSV2 or SMV1, a closely related virus
to SSV2 has been shown to be amenable to insertion of genetic material without loss of function.
Genome comparison between SMV1 and three related genomes revealed that some genes are not
present in all. For example, SMV3 has a much larger genome than SMV1, this suggests that large
DNA fragments could be inserted into the genome of SMV1 without loss of function. While the
potential for functionalization is there, the choice of modification strategy ultimately depends on the
application idea.
Throughout this work, focus has been on nanomedical applications but archaeal viruses also hold
great potential for applications in materials science. The natural characteristics of viruses such as
structural symmetry, uniformity, and potential for mutagenic alterations are unique properties for
materials, and have recently allowed the use of viruses in materials science and electronic
applications. For example, the rod-shaped filamentous bacteriophage M13 has been exploited as a
38
molecular template for ordering materials in a linear array which were used as seeds for subsequent
nanowire formation (106). Moreover, the piezoelectric and liquid-crystalline properties of M13
have been used for virus-based energy generation (107). Synthetic approaches in materials science
often use harsh conditions during either fabrication or use. As shown with SMV1, extremophilic
archaeal viruses are ideally suited for such harsh production schemes. Further, archaeal viruses
from extreme environments could expand the window of virus architectures available for materials
applications. The work presented here is only the starting point to the next exciting approaches that
may lead to new developments in nanobiotechnology using archaeal viruses as novel nanoplatforms
with applications in materials science and nanomedicine.
39
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I First insights into the entry mechanism of a spindle-­‐shaped extremophilic archaeal virus and its interaction with Sulfolobus host Kristine Buch Uldahl, Signe Bering Jensen, Yuvaraj Bhoobalan, and Xu Peng
Manuscript in preparation formatted as 4000 word research article
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46
First insights into the entry mechanism of a spindle-shaped
extremophilic archaeal virus and its interactions with
Sulfolobus host
Kristine B. Uldahl, Signe B. Jensen, Yuvaraj Bhoobalan, and Xu Peng# Danish Archaea Centre, Department of Biology, University of Copenhagen, Ole Maaløes vej 5, Copenhagen Biocenter, DK-­‐2200 Copenhagen N, Denmark Running title: Insights into the entry mechanism of SMV1 #Corresponding author E-­‐mail: [email protected] 47
Introduction
Archaeal viruses constitute an integral part of the virosphere and they are an ubiquitous feature of archaeal existence. Many Archaea are recognized as extremophiles, inhabiting extreme environments such as hot springs and solar salterns in high abundance (1, 2). Thus viruses infecting extremophilic Archaea can be considered key players in the complex population dynamics in these environments. Through host infection viruses can influence microbial diversity by introducing genetic diversity, affect host cell physiology and straight off kill their hosts by cell lysis (3, 4). Yet we only have a rudimentary understanding of archaeal virus-­‐host interactions. Much of the limited knowledge that we do have comes from studying the virus-­‐host interplay in Sulfolobales species. Of the less than 100 isolated archaeal viruses (ref), 19 infect hyperthermophilic Sulfolobus hosts. Among these viruses distinct characteristics are found; unique droplet-­‐ and spindle-­‐shapes, an exceptional pyramid-­‐like egress mechanism, and unique proteins with unknown functions (5). These distinctive characteristics are likely to influence the interplay with their hosts and give rise to unique life-­‐cycle traits. This has proven true for the rod-­‐shaped Sulfolobus virus, SIRV2, which has been shown to have an exceptional pyramid-­‐like egress mechanism. To date, SIRV2 remains the most well characterized archaeal virus and very little is known about the initial entry process and later egress mechanism of other archaeal viruses (6-­‐8). As a group, viruses with spindle-­‐shaped virions, single or two tailed are common in, and exclusive to the Archaea (9). Despite being the most common architecture found in Archaea-­‐
dominated habitats, studies of these viruses have mostly been confined to biochemical and genetic characterizations of their virions. Relatively little is known about the relationships between them and their hosts, in particular, insights into the entry process is lacking. Recently, it was suggested to group five of the large spindle-­‐shaped viruses together into a new superfamily based on structural similarities and a shared set of core genes (10). The group comprise of Acidianus two-­‐tailed virus (ATV), Sulfolobus tenchongensis spindle-­‐shaped virus 1 and 2 (STSV1 and 2), Acidianus tailed spindle-­‐shaped virus (ATSV), and Sulfolobus monocaudavirus 1 (SMV1). The virion structures of these large spindle-­‐shaped viruses are often pleomorphic and their tails also vary greatly in length. For example, ATSV tails range in length, between 35 and 720 nm, and ATV has been observed to develop two elongated tails once released from the host (10, 11). In order to elucidate the life-­‐cycles of these fascinating viruses a robust model system is needed. SMV1 was originally isolated from an acidic high temperature hot spring in Yellowstone National Park (12). Virions of SMV1 are spindle-­‐shaped (averaging 200 x 70 nm) with a single tail varying in length from 20 to 500 nm and a nose-­‐like structure on the opposite pole, which can extend to generate a second tail. SMV1 has a genome size of 48.8 kb with 51 putative ORFs (one major coat protein predicted). On infection in Sulfolobus islandicus Rey15A? growth retardation occurs but no evidence for cell lysis has been observed and no clear plaques has been seen on Gelrite plates (12). It has proven easy to reproduce SMV1 in S. islandicus to obtain high virus titers. Thus, this virus-­‐host system has been well established in our lab and represents a valuable model to study the virus-­‐host interactions in large spindle-­‐shaped viruses. Until now, the entry mechanisms of these viruses have not been thoroughly investigated. Here, we study the life cycle of SMV1 when infecting S. islandicus, with special focus on the early stages of infection. 48
Materials and methods
Virus propagation and purification
SMV1 was propagated in S. islandicus ΔC1C2 (13). The host culture was grown in Sulfolobus medium supplemented with 0.2 % (w/v) tryptone, 0.1 % (w/v) yeast extract, 0.2 % (w/v) sucrose and 0.002 % uracil (w/v) (TYS+U medium) (14). Cultures were started from -­‐80°C stock; cells were transferred to 50 mL TYS+U medium and incubated at 78°C. After 24 h of propagation, the 50 mL cell culture was transferred to 950 mL of pre-­‐heated (78°C) TYS+U medium. The culture was grown to an OD600 of 0.2-­‐0.3 (typically 24 h) at which time point the host culture was infected with SMV1 stock. The supernatant containing the virus particles were collected 48-­‐72 h post infection and concentrated by ultrafiltration using 1,000 kDa molecular-­‐weight cut-­‐off (MWCO) centrifugal filter units (Sartorius, Aubagne Cedex, France). The virus fraction was washed two times with 10 mL 10mM Tris AC buffer pH 6, to exchange the medium with storage buffer. The virus titer was determined by plaque assay as described below. Virus samples were stored at 4°C until used. Cultivation of Sulfolobus strains and growth conditions
5 Sulfolobales strains; S. islandicus LAL14/1 (15), S. islandicus HVE10/4 (16), S. islandicus REY 15A (16), S. islandicus ΔC1C2, S. solfataricus P25E6 (8) were tested for susceptibility to SMV1. All strains were cultivated in TYS+U medium at 78°C with aeration at 150 RPM. Six 50 mL flasks of each culture was set-­‐up; 3 x uninfected / 3 x infected with an MOI (multiplicity of infection) of 5. One millilitre was taken from each culture every 3 h for 48 h and OD600 values were measured. To investigate population dynamics, identical growth experiments were set-­‐
up with S. islandicus ΔC1C2 using an MOI range of 0.01-­‐5. The cultures were monitored for 48 h. For flow cytometry analysis, samples were taken at regular intervals from the infected (MOI 3) + control cultures and cells were fixed in 70% ethanol; 300 μL of culture was added to 700 μL 100 % ethanol and fixed for a minimum of 12 h at 4°C. One-step growth curve and plaque assay
Plaque assays were performed in order to establish a one-­‐step growth curve. Samples of cell-­‐
free supernatant from the time course infection of MOI 0.1 (see above) were serially diluted and 10 μL of each diluted sample was first mixed with 2 mL preheated fresh S. islandicus ΔC1C2 cells followed by mixing with 2 mL preheated 0.4% (w/v) phytagel. The mixture was layered over a 0.7 % (w/v) solid phytagel layer. Plates were incubated 2 d at 78°C. Single plaques were counted and plaque forming units (PFU) was determined. Virus stability experiments
All virus stability experiments were carried out using highly purified virus stocks. Treatments included (i) detergents (ii) proteases (iii) temperature (iv) UV radiation. Incubations were carried out in either Tris-­‐AC buffer pH 6 or treatment solution. After each treatment, infectivity was determined using the plaque assay, and virus particles were visualised by negative-­‐staining transmission electron microscopy. Briefly, virus particles were diluted 10-­‐
49
fold in the following treatment solutions; Triton X-­‐100 0.1% (w/v) or 0.01% (w/v), chloroform 5% (w/v), ethanol 5% (w/v), proteinase K (0.2 mg/mL), Tris AC buffer pH 1.5 or Tris AC buffer pH 6. The temperature conditions for the individual dilutions are listed in Table S1. To generate UV-­‐inactivated virus stocks, virus particles diluted 10-­‐fold in Tris AC buffer pH6, were irradiated in open petri dishes with either 40 mJ/cm2 or 1 J/cm2 of 254 nm UV light using a UV stratalinker 1800 (Stratagene, La Jolla, CA). Adsorption assay
For the adsorption assay, S. islandicus ΔC1C2 cells (OD600 = 0.2; 108 cells/mL) were infected using a MOI of 0.1. At defined time intervals, a sample of infected culture was removed and the adsorption stopped by immediate centrifugation (10,000 g, 5 min, room temperature (RT)). The number of remaining PFUs was determined by plaque assay and compared to the amount of virus present in a cell free control incubated at 78°C. The adsorption rate constant (k) was calculated using the formula, k = 2.3/(B)t x log10 P0/P (17) ,where P0 = PFU/mL at zero time, P = virus not adsorbed at time t min, (B) = concentration of host cells as number of cells/mL. Membrane integrity test
The membrane integrity of host cells was assessed using the Live/Dead BacLight bacterial viability kit (Molecular Probes, OR, USA) with minor modifications to the manufacturer’s protocol. Briefly, 500 µl samples were taken at regular intervals from the infected (MOI 3) + control cultures and cells were collected by centrifugation (1,000 g, 20 min, RT). The cells were re-­‐suspended in 1 mL incubation medium containing 0.3 % (w/v) ammonium sulphate, 0.05 % (w/v) potassium sulphate, 0.01 % (w/v) potassium chloride, and 0.07 % (w/v) glycine, pH ∼5.2. The samples were centrifuged again under same conditions. Pellets were re-­‐
suspended in 200 μL incubation medium containing SYTO9 (0.0068 mM) and Propidium Iodide (0.04 mM) for 15 min at RT. Samples were analysed immediately after staining in an ApogeeFlow A-­‐40 flow cytometer illuminating with a 488 nm laser. ®
Flow cytometry
Samples were fixed in 70 % ethanol (v/v) and stored at 4°C. When all the samples were collected the fixed cells were centrifuged at 3,000 g for 20 min and re-­‐suspended in 1 mL Tris MgCl2 buffer containing 10 mM Tris (pH 7.5) and 10 mM MgCl2. The samples were centrifuged again under same conditions. Pellets were re-­‐supended in 150 μL fresh staining solution containing mithramycin (100 μg/mL) and ethidium bromide (20 μg/mL) for 1 h. At all steps, the samples were kept cold. Samples were analyzed in an ApogeeFlow A-­‐40 flow cytometer (Apogee Flow Systems, Hemel Hempstead, UK) illuminating with a 405nm laser. Interaction of SMV1 with S. islandicus ΔC1C2 cells
SMV1 virions were incubated with S. islandicus ΔC1C2 cells (high MOI) in a thermoblock (Fisher Scientific) at 78°C for 10 min, the tube was inverted several times to prevent pellet formation. Immediately after incubation the sample was prepared for TEM. 50
TEM
Virus containing fractions were adsorbed onto carbon-­‐coated copper grids for 5 min and stained with 2% (w/v) uranyl acetate. Images were recorded using a JEM-­‐1010 transmission electron microscope (JEOL, Tokyo, Japan) with a Gatan digital camera 792. Results and discussion
SMV1 exhibits a distinct host-­‐range profile even among closely-­‐related Sulfolobales strains. To find a suitable host for studying the life cycle of SMV1, we infected a range of well-­‐
established lab strains of Sulfolobales to test for susceptibility. Previous studies have shown that SMV1 susceptible cultures show clear growth retardation compared to non-­‐susceptible cultures. Infection studies showed S. islandicus REY15/4 and the deletion mutant (S. islandicus ΔC1C2) to be highly susceptible to SMV1 infection. Growth of the infected strains was strongly inhibited at a multiplicity of infection (MOI) of 5. REY15/4 displayed observable growth retardation 14 hours post infection (hpi) whereas ΔC1C2 displayed growth retardation already 6 hpi. The uninfected control cultures showed normal growth with a generation time of about 8 h (Fig. 1A). The ΔC1C2 strain was derived from REY15/4 carrying a ~120 kbp deletion. The deleted region includes the type I-­‐A CRISPR system, the two repeat-­‐spacer arrays, some toxin-­‐antitoxin genes and hypothetical genes, which likely confers some immunity to SMV1. Growing cultures of S. islandicus HVE10 and S. islandicus LAL 14/1 showed no indication of susceptibility to SMV1 with growth curves identical for infected and non-­‐infected cultures (Fig. S1). Interestingly, cultures of S. solfataricus P2 5E6 show growth retardation between 14 – 23 hpi at which point the cultures start growing again with a growth rate comparable to the control (Fig. S1). SMV1 has been confirmed by plaque assay to reproduce in P2 5E6 (data not included). Investigation of the recovery of P2 5E6 was not pursued in this work but could prove interesting for future studies investigating if SMV1 has host-­‐dependent life-­‐cycle traits. Based on the results we selected ΔC1C2 as the host for further investigations into viral life-­‐
cycle traits of SMV1. First, we infected ΔC1C2 cultures at a range of MOIs (0.01 – 5) to observe for differences in growth retardation. The MOIs were calculated based on viral titers enumerated as PFU/mL. Figure 2 clearly shows that the higher the MOI the sooner the growth retardation occurs, at MOI 5 (∼99 % of the cells are infected) growth retardation is observed at 6 hpi compared to 12 hpi at MOI 0.01 (∼1 % of the cells are infected). A one-­‐step growth curve revealed a dramatic increase of extracellular virus titer at around 8 hpi (Fig. 2A), which is comparable to the release time of STSV1 (18) as well as to that of SIRV2 (8). This correlates with the observed delayed growth retardation at MOI 0.01; at 12 hpi new viral progeny from the first replication cycle would have been released and infected the remaining uninfected cells, at which point the growth of the whole culture would be affected. To gain insights into the initial stages of SMV1 entry, we followed the kinetics of SMV1 adsorption to ΔC1C2 cells. The adsorption was very efficient with 50 % of virions being bound to cells within 1 min of infection (Fig. 2B). Further incubation of the virus in presence of the host cells resulted in additional virion binding; ∼80 % of virions were bound within 20 to 30 min post infection (pi). All adsorption assays were conducted under optimal growing 51
conditions for the ΔC1C2 host cells, i.e. at 78°C and pH 3.5. To ensure that the observed loss of virus titer were not due to high temperature and/or acidic effects but indeed could be assigned to virus adsorption, we performed a cell-­‐free control in which the same amount of SMV1 as used for the infection was added to ΔC1C2 growth medium. The virus titer of the control did not change over the 30 min of incubation (conditions as above). The adsorption rate (calculated as 7 x 10-­‐9 mL min-­‐1 at 1 min pi) is very rapid and comparable to the even faster adsorption rate observed for SIRV2 (6). SMV1 and SIRV2 are the only two hyperthermophilic archaeal viruses for which the adsorption rate is known. The only other group of archaeal viruses for which adsorption rates has been studied are the viruses of halophilic Archaea, which often bind to their hosts extremely slow e.g. 30 % adsorption over 3 h is observed for His1 (19, 20). The rapid adsorption rates of hyperthermophilic viruses are hypothesized to minimize the time they spent in the hostile extracellular environment with boiling temperatures and acidic pH (6). Almost nothing is known about the viral entry mechanisms of archaeal viruses. Studies of viruses infecting the other domains of life have observed three common entry mechanisms; penetration (non-­‐enveloped), membrane fusion (enveloped), and endocytosis (enveloped) (21). SMV1 has previously been shown to have lipids at the outer capsid concurrent with an enveloped virus (ref). Other spindle-­‐shaped viruses such as SSV1 and SSV2 have well-­‐
established lipid envelopes. To elucidate the possible entry mechanism of spindle-­‐shaped archaeal viruses, we followed the interaction of SMV1 and ΔC1C2 cells by transmission electron microscopy (TEM) just after infection. In general, we observed three to four virus-­‐cell conformations (Fig. S2) which led us to propose a fusion entry mechanism for SMV1 comprising of three distinct stages. The first stage is the initial attachment to the host cell; the attachment occurs at the nose-­‐like end and from the TEM images the receptor appears to be highly abundant as > 30 virions can be attached at the same time. The second stage, we refer to as the alignment stage. It appears the virion align along the host cell, initially the virion body are adsorbed along the host S-­‐layer while the tail/tails are still unabsorbed. At some point the whole virion including tails are absorbed along the S-­‐layer and the virus particle is coating the cell. The final stage is the fusion stage where the virion appears to be flattened against the S-­‐layer and the virus morphology “disappears” (Fig. 3). Further studies are needed to clearly establish the fusion mechanism and other spindle-­‐shaped archaeal viruses should be investigated to see if fusion is a common entry mechanism for spindle-­‐shaped archaeal viruses. Upon SMV1 infection clear growth retardation is observed in ΔC1C2 cells, however, it is less clear whether virus release lyses the cells or leaves them unable to recover thus preventing further growth. To investigate whether SMV1 has a lysogenic or lytic life-­‐cycle, we performed a detailed cellular study of the infection cycle of SMV1. We investigated chromosome degradation and membrane disruption, using flow cytometry time-­‐course analysis in combination with Live/Dead® baclight viability detection. The cell size and intracellular DNA content in uninfected and SMV1-­‐infected cultures (MOI 3) over time were monitored by flow cytometry. The DNA content distributions of the control cultures (Fig. 4, left) were typical for exponentially growing Sulfolobus cells (22), with a majority of the cells containing 2 chromosomes. In the infected cultures (Fig. 4, right) cells with a very high DNA content (> 2 genomes) started to appear already 4 hpi and continued to increase in proportion till 22 hpi whereas the proportion of cells containing 1-­‐2 genome equivalents decreased. At 16 and 22 52
hpi a large majority of the cell population had DNA content above the detection range of the assay (>> 2 genomes). After extruding from the host cell, SMV1 particles often remain attached to the surface of the cell, and layers of virus particles have occasionally been observed to surround a single or multiple host cells (Fig. S3A). A similar tendency has been observed for STSV1 and host cells (18). Thus giving rise to particle aggregates with very high DNA content, which are outside of the 1-­‐4 genome size distribution range of the assay. During the 22 h experiment, equivalent to two viral replication cycles, no significant chromosome degradation occurred in the SMV1-­‐infected cultures. Moreover, the Live/Dead® baclight assay only showed a negligible increase in the amount of ‘dead’ cells in the SMV1-­‐infected culture over 48 h compared to the control culture (Fig. S3B). Thus, no indications of cell lysis were observed and SMV1 appears to be nonlytic like the related STSV1 and the spindle-­‐
shaped fuselloviruses. SMV1 particles isolated and purified immediately after release appear spindle-­‐shaped with no visible tails (Fig. 5A). Upon release and if left in the cell culture at 78°C, the virions start developing tails; sometimes tail development occurs at both poles with one tail being shorter than the other. Purified SMV1 particles are often observed in rosette-­‐like structures (Fig. 5B). Together this gives rise to a very heterogenic virus population as seen in figure 5C and 5D. The closely related ATV virus displays similar extracellular tail development independent of its host (23). The ability of SMV1 and ATV to undergo major extracellular morphological development is a unique feature of these two archaeal viruses, not observed elsewhere in the virosphere. The reason for the extracellular development of the virions is still unclear but their exceptional development may constitute a strategy for survival under extreme conditions in an environment with low cell density (23). Also, based on the observations of the fusion entry mechanism, we hypothesize that the tails play a key element in fusion with the host cell during the alignment phase. Other extremophilic archaeal viruses have shown remarkable stability under a range of conditions and recent studies have explored SMV1 as a potential nanoplatform (ref). Therefore, we wanted to assess the tolerance of SMV1 particles towards natural and non-­‐
natural harsh conditions. Highly purified virions were treated with different detergents, solvents and proteinase K as well as a variety of pH, temperature, and UV conditions. After each treatment, viral infectivity was determined and the appearance of the virus particles was assayed by negative stain TEM. Overall, SMV1 particles demonstrated extreme stability under a range of conditions; freezing without cryoprotectants and suspension in 5 % v/w ethanol had no effect on infectivity and appearance, the other treatments inactivated the virus particles to various degrees. However, TEM observations indicated that the only conditions degenerative to SMV1 structural integrity involved autoclaving conditions and pH <1.5 (Table S1). Some of the conditions that inactivated SMV1 resulted in a transformation of the spindle-­‐
shaped virion body, especially proteinase K and boiling conditions triggered the development of one or two tails at the poles of the virion body. It is likely that the proteinase treatment of SMV1 particles interacted with the capsid proteins, leading to the observed conformational changes. Tail development under boiling conditions is concurrent with the observations of tail development at temperatures close to that of the natural habitat. Thus indicating high temperature to be a very likely environmental trigger for virion transformation. 53
Hot springs, the natural habitat of most spindle-­‐shaped viruses receive a high amount of natural solar irradiance (25) therefore we wanted to determine the inherent resilience of SMV1 exposed to UV irradiance. We used two different UV doses to test for inactivation of SMV1; a very high dose (1 J/cm2) and the European standard (40 mJ/cm2) based on the inactivation of most pathogens (26). With the lower dose ∼98 % of the virus particles were inactivated and at the higher dose 100 % inactivation was observed by using plaque assays. To test the complete inactivation of SMV1 at the higher UV dose, we added 100 µl of the inactivated virus suspension to a growing culture of ΔC1C2 cells and performed a one-­‐step growth curve experiment to see if we could observe virus reproduction over time. Interestingly, after 6 h we started observing virus plaques and after 8 – 10 h a virus titer of 103 PFU/mL was observed by plaque assay, indicating that the initially added virus had reproduced (data not included). Thus, a complete inactivation had not occurred even at the very high UV dose. The tendency of SMV1 to form rosette-­‐like structures outside of the host probably provides a certain amount of protection against UV exposure; the rosette-­‐like aggregates shield a small percentage of the virus particles from direct UV light, thus a subpopulation of viruses remain infectious even under extreme irradiation. Rosette-­‐like aggregates have also been observed for the hyperthermophilic fuselloviruses, where it appears thin protruding fibers at one pole can attach to the same fibers in other viruses leading to the characteristic rosette-­‐like structures. Similar fibers have been observed at one tail end of SMV1 (Fig. S4). We hypothesize that the ability of spindle-­‐shaped viruses to form protective rosette-­‐like aggregates is a natural defence mechanism against harsh environmental conditions and is one of the reasons for their widespread success in Archaea dominated habitats (27-­‐29). In addition, the UV exposure experiment makes us very cautious when interpreting infectivity by plaque assay. If no infectivity is seen by plaque assay it should not be interpreted as an absolute but be followed-­‐up by a one-­‐step growth curve analysis to see if the virus reproduces to observable numbers. Finally, we want to stress the difficulties in estimating precise titres of infectious virus particles. As seen in Fig. 3 more than 30 virus particles can attach to one cell but will give rise to only one plaque. Thus, plaque assays are far from a precise enumeration method. Also, we have used a nanoparticle analyser to estimate the nanoparticle count in a highly purified virus suspension, which gave a 10-­‐fold higher particle count than by plaque assay. Another study enumerated the spindle-­‐shaped virus SSV9 by plaque assay and qPCR and similarly found a 10-­‐fold difference (30). Suggesting this to be a general observation for spindle-­‐shaped viruses. Concluding remarks Our study provides valuable insight into the entry mechanism of one of the large spindle-­‐
shaped archaeal viruses, SMV1. Our observations suggest that SMV1 virions enter into host cells via a fusion entry mechanism, involving three distinct stages; attachment, alignment, and fusion. Interestingly, we observed more than 30 virions attached to the surface of a single host cell, indicating a relative abundance of viral receptors. Upon infection, the replication of the virus retarded host growth but did not cause lysis of the host cells. As with ATV, SMV1 shows remarkable virion plasticity by developing a tail/tails independent of the host. An extraordinary feature only observed for these two hyperthermophilic spindle-­‐shaped viruses. In our stability experiments, SMV1 particles demonstrate an exceptional resilience to a range of harsh conditions, especially at extremes of temperature and UV irradiance. We hypothesize 54
that the rosette-­‐like aggregates observed for SMV1 and other spindle-­‐shaped viruses act as a natural defence mechanism, thus increasing their resilience when exposed to harsh environmental conditions and is one of the reasons for their widespread success in Archaea dominated habitats. The work presented here is only the starting point for future studies into the viral traits of the large spindle-­‐shaped archaeal viruses. Funding information
This research received no specific grant from any funding agency in the public, commercial, or not-­‐for-­‐profit sectors.
Acknowledgments
We thank members of the Danish Archaea Centre, Copenhagen, for stimulating discussions and helpful advice. 55
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Figures Figure 1 (A) Suceptibility of S. islandicus REY 15/A and S. islandicus C1C2 to SMV1 infection. OD600 was measured over a time period of 2 days in cultures with (open symbols) or without (filled symbols) the addition of SMV1 (MOI 5). Cultures were incubated in triplicates at 78°C. (B) Growth inhibition of S. islandicus C1C2 to SMV1 infection at different MOIs (range 0.01 – 5). 58
Figure 2 (A) One-­‐step growth curve of SMV1 infection of S. Islandicus C1C2. SMV1 was added at an MOI of 0.1. The plaque forming units (PFU) are plotted against time. (B) Kinetics of SMV1 adsorption to cells of S. Islandicus C1C2. Cells were infected with SMV1 using an MOI of 0.1 at 78°C. The number of unbound virus particles was determined at different time points post infection as described in Materials and Methods. 59
Figure 3 Electron micrographs of SMV1 interaction with S. islandicus ΔC1C2 cells. Samples were collected 10 min post infection and negatively stained for TEM. Scale bar left 1μM, scale bars right 200 nm. The observed interactions led to a proposed fusion entry mechanism for SMV1. The drawings below the electron micrographs illustrates the fusion mechanism divided into three distinct stages; attachment, alignment, and fusion. 60
Figure 4 Flow cytometry time course analysis of S. islandicus ΔC1C2 cells infected by SMV1. Left panel presents cell size and DNA content distribution from an uninfected culture. Right panel presents cell size and DNA distribution from a culture infected with SMV1 (MOI ≈ 3). The virus was added just before time point 0 h. 61
Figure 5 Electron micrographs of different forms of Sulfolobus monocaudavirus, or SMV1. SMV1 particles isolated immediately after release occur tailless: a. SMV1 develops 1 or 2 tails outside of the host cell: b-­‐d,. The development occurs at high temperatures >75°C. Often one longer and one shorter tail is observed (blue arrows). Only one pole appears to have short tail fibres which can attach to other tail fibres to form characteristic rosettes: b. All preparations were negatively stained with 2% uranyl acetate. 62
Supporting information Figure S1 Susceptibility of S. Sulfolobus P2 5E6, S. Islandicus LAL, and S. islandicus HVE10 to SMV1 infection. OD600 was measured over a time period of 2 days in cultures with (open symbols) or without (filled symbols) the addition of SMV1 (MOI 5). Cultures were incubated in triplicates at 78°C. 63
Figure S2 Electron micrographs of SMV1 infected S. islandicus ΔC1C2 cells at 10 min post infection. The alignment stage of viral infection is shown with purple arrows. All preparations were negatively stained with 2% uranyl acetate. 64
Figure S3 A) Electron micrograph of a S. Islandicus ΔC1C2 culture infected with SMV1 (high MOI) 22 h post infection. Scale bar 1 μM. B) A time course analysis of the percentage of live (light grey) and dead (dark grey) cells in a S. islandicus ΔC1C2 culture infected with SMV1 (MOI ≈ 3) as compared to an uninfected control. Figure S4 Electron micrograph of negatively stained SMV1 particles. Blue circles indicate observed tail fibers protruding from one tail end of the virion body. 65
Table S1 Stability experiments 66
67
68
II Recognition of extremophilic archaeal viruses by eukaryotic cells; an emerging nanoplatform from the Third Domain of Life Kristine Buch Uldahl, LinPing Wu, Arnaldur Hall, Xu Peng, and Moein Moghimi
Manuscript submitted to Biomaterials
69
70
Recognition of extremophilic archaeal viruses by eukaryotic cells: an
emerging nanoplatform from the third domain of life
Kristine B. Uldahl1,†, Linping Wu2,†, Arnaldur Hall2, Xu Peng1,*, and S. Moein Moghimi2,3,*
1
Danish Archaea Centre, University of Copenhagen, DK-2200 Copenhagen N, Denmark
2
Nanomedicine Research Group, Centre for Pharmaceutical Nanotechnology and Nanotoxicology,
Department of Pharmacy, University of Copenhagen, Universitetsparken 2, DK-2100 Copenhagen
Ø, Denmark
3
NanoScience Centre, University of Copenhagen, Universitetsparken 5, DK-2100 Copenhagen Ø,
Denmark
†
Contributed equally.
*Corresponding authors: [email protected] / + 45 35322018 (XP); [email protected] / +
45 35336528 (SMM)
71
Abstract
Archaeal viruses belong to an abundant category of viruses from the third domain of life with
specializations that allow their survival in extreme environments. Here, we selected two archaeal
viruses Sulfolobus monocaudavirus 1 (SMV1) and Sulfolobus spindle shaped virus 2 (SSV2) owing
to their unique shape, hyperthermostable and acid-resistant nature and studied their interaction with
mammalian cells. Accordingly, we followed viral uptake, intracellular trafficking and cell viability
in human endothelial cells of brain (hCMEC/D3 cells) and umbilical vein (HUVEC) origin. Unlike
SMV1, SSV2 differentiates between HUVECs and hCMEC/D3 cells, thus opening a path for
selective cell targeting. On internalization, both viruses localize only to the lysosomal
compartments. Neither SMV1, nor SSV2 induced any detrimental effect on plasma membrane and
mitochondrial oxidative phosphorylation. We discuss the potential application of archaeal viruses in
bioengineering and future development of multifunctional vectors for cellular manipulation and
sensing, since these species have never been reported to integrate into human or any other
eukaryotic genomes.
Keywords: Archaeal viruses; Endothelial cells; Sulfolobus monocaudavirus 1; Sulfolobus spindle
shaped virus 2; Nanoplatforms; Viral vectors
72
1. Introduction
Viruses are receiving increasing attention as novel nanoplatforms with applications in
materials science and medicine (1). Viruses demonstrate remarkable features including plasticity,
coordinated assembly, and site-specific delivery of nucleic acids. Viruses are also amenable to
genetic engineering, their internal cavity may be filled with therapeutic agents, and the functional
groups on the virus capsid may be modified with biomolecules, synthetic polymers and diagnostic
agents (2). Accordingly, viruses could provide basis for the development of alternative
multifunctional vectors and theranostic platforms (3). Within such notions, plant viruses and
bacteriophages receive special attention, as they are considered non-infectious and non-hazardous
in humans. Another group of viruses that fits this criterion is archaeal viruses, a highly diverse and
abundant category of viruses from the third domain of life [4], yet their potential remains untapped.
Archaeal viruses offer an ideal search pool for novel nanoplatforms as they have several attractive
features. They are non-pathogenic, offer unique morphologies, and have specializations to survive
in extreme environments [5]. All known archaeal viruses infect extremophilic Archaea, and are thus
adapted to survive the harsh environments of the host, making them extremely stable entities [4,5].
As a group, archaeal viruses show distinct morphologies not found in bacteriophages or plant
viruses. These include lemon-, bottle-, and droplet-shape (4). Accordingly, due to their unique
shape and inherent properties, archaeal viruses may prove as interesting vehicles for differential
targeting of eukaryotic cells. Furthermore, size and shape have been identified as key factors
influencing circulation half-life, biodistribution and cellular uptake of particulate drug delivery
vehicles [6]. Although several articles suggest archaeal viruses as promising nanoplatforms (5-7), to
the best of our knowledge there are no reported studies investigating the uptake and intracellular
fate of archaeal virus in eukaryotic cells, which is a first step in evaluating their potential as a
nanoplatform for cellular targeting and manipulation.
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Here, we studied two archaeal viruses; Sulfolobus monocaudavirus 1 (SMV1) and
Sulfolobus spindle shaped virus 2 (SSV2) as candidate nanoplatforms. Both viruses infect hosts
from the archaeal family Sulfolobaceae. These organisms are found in volcanic hot springs, and are
considered hyperthermophilic acidophiles with optimal growth at 80 °C and pH 2-3 (8). The
fusellovirus SSV2 is a lemon-shaped dsDNA virus (with a genome size of 14.8kb) of 65-70 nm in
size bearing short flexous tail fibers at one pole. SMV1 share many of the same characteristics, but
it is slightly larger (120 nm) with a genome size of 48.8kb (9). SMV1 and SSV2 were selected
owing to their unique shape, hyperthermostable and acid-resistant nature. Furthermore, both species
are well-established laboratory strains with the potential for up scaling. We have investigated the
uptake, intracellular fate, and safety of fluorescently labelled SMV1 and SSV2 in two different
endothelial cell types of human origin, hCMEC/D3 and HUVEC, providing the first insights into
the interaction between archaeal viruses and eukaryotic cells.
2. Materials and methods
2.1. Production and purification of virus particles
SSV2 was propagated in S. solfataricus strain 5E6, a highly susceptible strain as described
previously (10). SMV1 was propagated in S. islandicus CRISPR deletion mutant delta C1C2 (11).
Both host cultures were grown in Sulfolobus medium supplemented with 0.2% (w/v) trypton, 0.1%
(w/v) yeast extract, 0.2% (w/v) sucrose and 0.002% (w/v) uracil (TYS+U medium) (12). Cultures
were started from -80°C stock; cells were transferred to 50 mL TYS+U medium and incubated at
78°C. After 24h of propagation, the cell culture was transferred to 950 mL of pre-heated (78°C)
TYS+U medium. The culture was grown to an optical density at 600 nm (OD600) between 0.2-0.3
(typically 24 h) at which time-point the host culture was infected with virus isolate. The
supernatants containing the virus particles were collected 48-72 h post-infection and concentrated
74
by ultrafiltration using 1,000 kDa molecular-weight cut-off (MWCO) centrifugal filter units
(Sartorius, Aubagne Cedex, France). Additionally, the virus particles were purified by
ultracentrifugation through a 10-40% (w/v) continuous Iodixanol gradient. Continuous gradients
were prepared by sequentially layering 10, 20, 30 and 40% (w/v) Iodixanol solution (OptiPrep™,
Axis-Shield PoC AS, Oslo, Norway) in 10 mM Tris-HCl, pH 6.0 into 14 mL centrifuge tubes
(Beckman Coulter UK Ltd., High Whickham, Bucks, UK). The gradients were left in the dark at
4°C overnight and then ultrafiltrated virus preparations were layered over the continuous gradient (1.0 mL) and centrifuged in a SW-41 rotor (Beckman Coulter) for 6 h at 95,000 g at 4°C.
An opaque virus band was visible and recovered and an additional ultrafiltration step was
performed. The virus fraction was washed three times with 10 mL of 10 mM Tris-HCl, pH 6.0 to
remove Iodixanol and to prevent interference with downstream processes. Transmission electron
microscopy (TEM) was used to confirm the presence of viral particles in the recovered fractions,
and the number of plaque forming units (PFU)/mL was calculated (see below). The virus
preparations were then stored at 4°C until used.
2.2. Virus labelling
For labelling of viruses, 1.0 µL of PKH67 dye (Sigma, St. Louis, MO) [15] was dissolved in
1.0 mL of Diluent C (Sigma, St. Louis, MO) immediately before the start of procedures. Two
volumes of diluted PKH67 was added to one volume of viral suspension (250.0 µL), and mixed by
pipetting. After 3 min, the reaction was stopped by adding three volumes of 1% (w/v) BSA, then
incubated for 1 min to allow binding of excess dye. The suspension was then centrifuged at 20,000
g for 20 min at 4°C and the supernatant were discarded. Labelled virus were re-suspended in 3 mL
of 10 mM Tris-HCl, pH 6.0 and centrifuged at 20,000 g for 20 min. The washing step was repeated
and the final pellet were re-suspended in 250.0 µL Tris-HCl and stored at 4°C in the dark.
75
2.3. Determination of virus size and concentration
Nanoparticle Tracking Analysis (NTA) was used for determination of viral hydrodynamic
size distribution and concentration before and after labelling [16]. Briefly, samples were diluted
x106 with 250.0 µL Tris-HCl and monitored with an LM20 NanoSight mounted with a blue (405
nm) laser (Nanosight, Amesbury, UK). Data analysis was performed using the NanoSight 2.3
software [16]. All measurements were performed in triplicate and at room temperature.
2.4. Transmission electron microscopy
Viral preparations were placed on a carbon-coated copper grid for 5 min, and stained with
2.0% (w/v) uranyl acetate. Images were recorded using a JEM-1010 transmission electron
microscope (JEOL, Tokyo, Japan) with a Gatan digital camera 792.
2.5. Plaque assay
The approximate virus titer before and after fluorescent labelling was determined by plaque
assays. Serial dilutions (10.0 µL) of viral preparations were mixed with a sample (2.0 mL) of the
exponentially growing host culture (2 x 108 cells); S. islandicus ΔC1C2 for SMV1 infection and S.
solfataricus 5E6 for SSV2 infection, respectively. The mixture was incubated for 30 min at 78°C to
allow the adsorption of the virus to the host cells. Immediately following the addition of 2 mL of
TYS+U medium containing 0.25% (w/v) Gelrite (78°C), the sample was layered over a premade
0.8% (w/v) Gelrite plate (78°C). The plates were incubated for 2 days at 78°C. SMV1 plaques
appeared as small clear halos, whereas SSV2 plaques appeared excessively turbid making them
difficult to detect and enumerate.
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2.6. Imaging of infected host cells
From an exponentially growing S. islandicus ΔC1C2 culture (OD600 ∼ 0.2), 5.0 mL were
collected and incubated with SMV1 purified stock at an MOI of 10. At 5 and 30 min post infection
(mpi) 1.0 mL cells were pelleted at 8,000 g for 5 min and then re-suspended in 500 µL 4% (v/v)
formaldehyde. After 10 min incubation at room temperature, cells were pelleted at 8,000 g for 3
min then washed with PBS buffer, pH 7.4. To permeabilize cells, pellets were suspended in 500 µL
PBS containing 0.1% (v/v) Triton X-100 and incubated at room temperature for 15 min. Cells were
then washed three times with PBS to thoroughly remove all traces of the detergent. Pellets were resuspended in 100-200 µL PBS, and 10.0 µL were spread onto poly-D-lysine coated coverslips. After
air-drying, the coverslips were washed with PBS then stained with DAPI (10.0 µM) for 15 min at
room temperature. After staining cells were washed three times with PBS, a cover glass was placed
on top and the edges were sealed with clear nail polish. Coverslips were stored in the dark at 4°C.
Differential interference contrast and fluorescent images were acquired in Volocity (Improvision)
with a cooled Orca-ER CCD camera (Hamamatsu) mounted on a Zeiss AxioImager Z1 microscope
(Carl Zeiss, Germany). All images were captured at 100-fold magnification using a PlanApochromat 100×, 1.4 NA objective lens.
2.7. Cell uptake studies
Human brain capillary endothelial cell line hCMEC/D3, transduced by lentiviral vectors
incorporating human telomerase or SV40 T antigen, were grown in EBM-2 medium supplemented
with 5.0% (v/v) FBS, hydrocortisone (1.4 × 10−6 M), basic fibroblast growth factor (1 ng/mL), 1.0%
(w/v) penstrep and 10 × 10−9 M HEPES [16]. Human umbilical vein endothelial cells (HUVECs)
were grown in DMEM with 20.0% (v/v) FBS on gelatin-coated flasks. All cells were maintained at
37oC in a humidified atmosphere (air supplemented with 5% CO2). For the uptake study,
hCMEC/D3 cells and HUVEC cells (2 × 104 /cm2) were seeded on 24-well plates (Corning, USA)
77
and grown for 24 h at 37°C, and 5% CO2 to 60%–70% confluence. Afterwards, cells were washed
three times with pre-warmed PBS and the uptake studies were initiated by adding different
concentration of PKH67 labelled viral particles. After viral challenge, the medium was removed
and cells were washed three times with pre-warmed PBS and harvested by trypsinization. A total of
10,000 cells, suspended in ice-cold PBS, were analyzed by flow cytometry (FACS Array Cell
Analysis, BD, USA). For live-cell microscopy and organelle tracking, cells were seeded on eightwell Lab-Tek chamber slides (Nunc, Naperville, IL) for 1 day and then labelled with CellLight
Reagents BacMam 2.0 Early endosomes-GFP, Lysosomes-GFP, Golgi-GFP, Endoplasmic
reticulum (ER)-GFP, and Tubulin-GFP in accordance with the manufacturer’s protocol. Live cell
imaging was performed on a Leica AF6000LX microscope equipped with a 63× (numerical
aperture 1.47) oil objective using 1.6× magnification and analyzed (13). The co-localization
analysis was processed with Image J to calculate Manders’ coefficient as described previously (14).
2.8. Cell functionality and viability tests
Lactate dehydrogenase (LDH) release, and cell respiration was investigated as described in
detail elsewhere [16,17]. LDH release from hCMEC/D3 cells and HUVECs was measured at 24 h
post incubation with viruses using CytoTox96 Non-Radioactive Cytotoxicity Assay kit (Promega).
To investigate the possible effect of virus on cell respiration, cells were seeded in XF96 V3 cell
culture microplates (Seahorse Bioscience) at 1.0 × 104 cells/well in growth medium for 24 h.
Following viral challenge at 37°C and 5% CO2 for 24 h, oxygen consumption rate (OCR) was
monitored in real-time using XF96 Analyzer (Seahorse Bioscience) to measure different respiratory
states, and to calculate the coupling efficiency of mitochondrial oxidative phosphorylation
(OXPHOS) and the respiratory control ratio (RCR) [17,18].
78
3. Results and discussion
3.1. Viral morphology and size distribution
Morphological characteristics of SMV1 and SSV2 were studied by TEM and representative
micrographs are presented in Figure 1. SMV1 is lemon-shaped with an extending tail region at one
end. SMV1 appears both individually and in a rosette-like arrangement, where virus-virus
interaction appears to be mediated through the tail regions. Although, the general morphology of
SMV1 remains unaltered after PKH67 labelling, a fluorescent membrane dye that efficiently labels
enveloped viruses non-covalently [15], some virions appear more electron dense. SSV2 is also
lemon-shaped bearing flexous tail fibres at one pole, but these viruses are smaller than SMV1
(Figure 1). This species also form rosette-like structures of 3-6 virions as seen in TEM (Figure 1).
Again, their morphological characteristics appear to be preserved on PKH67 labelling and with
some increase in electron density.
NTA was used for sizing and counting of viral preparations.
NTA deduces the size on a virus-by-virus basis, relating the changing position of scattering viruses
under Brownian motion [19]. NTA, however, assumes sphere equivalent hydrodynamic diameter
particle size. Accordingly, SMV1 displays a narrow size distribution with a mean sphere equivalent
hydrodynamic diameter of 114 ± 2 nm (n = 3) and 120 ± 1 nm (n = 3), with corresponding modes of
108 ± 2 nm and 120 ± 3 nm, before and after labelling, respectively (Figure 1). NTA analysis
further reveals existence of a small population of ≤50 nm particles, presumably arising from
particulate contaminants and debris that could not be removed during the purification steps. This
population, however, seems unaffected after labelling and still present. Another interesting feature
of the NTA analysis is the slight increase in the population of ≥150 nm species after PKH67
labeling, which most likely is a reflection of aggregate formation among particles in the 75-100 nm
ranges, since their concentration drops compared with untreated viruses. These differences are
further reflected in scatter plots of relative intensity versus particle size, which allow differences in
79
particle composition to be explored. The results reveal broad scattering of particles in 50-150 nm
size ranges with relative intensities of 1-10 arbitrary units (au), whereas larger particles/aggregates
predominantly scatter with relative intensities <5 au (Figure 1). PKH67-labeling causes
considerable drop in the proportion of high scattering (>5 au) SMV1 particles, and concomitantly
increases the proportion of particles that scatter <5 au, including particles ≥150 nm in size.
NTA determination also shows a narrow size distribution for SSV2 with a mean sphere
equivalent hydrodynamic diameter of 76 ± 1 nm (n = 3), and 75 ± 2 nm (n = 3), with corresponding
modes of 63 ± 5 nm, and 65 ± 4 nm, before and after PKH67 labelling, respectively. Again, a small
proportion of small particles (20-30 nm) are observable before and after labelling, which may
represent the presence of some impurities (Figure 1). Unlike SMV1 preparation, PKH67 did not
increase the proportion of larger particles (>100 nm). Considering the hydrodynamic diameter of
SSV2, the detection of a minor population of ≥150 nm may represent SSV2 rosettes that were
observed in the electron micrographs. Finally, the scatter-plots show that the bulk of SSV2 particles
scatter with intensities ≤1 au, but after labelling a higher proportion of SSV2 scatter >1 au (Figure
1).
3.2. Viral infectivity after PKH67 labelling
Infectivity was measured by plaque assay with host cells. The results in Figure 2 confirm
that both viruses remain infectious after PKH67 labelling. However, SSV2 plaques appeared
excessively turbid thus making them difficult to enumerate. Accordingly, the infectivity of SSV2
must be viewed cautiously. Next, we infected a culture of S. islandicus ΔC1C2 with PKH67
labelled SMV1 to investigate whether PKH67 signal colocalize with the host cells stained with
DAPI. Co-localization analysis revealed a Pearson’s correlation coefficient (PCC) of ∼ 0.9. Colocalization coefficients M1 and M2 demonstrated that the majority of the green pixels (deriving
80
from PKH67) co-localize with the blue DAPI pixels, whereas ~ 65% of the blue pixels (host cells)
co-localize with the green pixels (Figure 2). These observations suggest that at least the majority of
the PKH67 signal is derived from the labelled SMV1, which remains infectious. Finally, we used S.
acidocaldarius, which is not a host to any known archaeal viruses, to confirm selective infectivity.
Indeed, no co-localizing signals were observed with DAPI stained S. acidocaldarius cells incubated
with PKH67-labeled SMV1 under the same condition as used for S. islandicus ΔC1C2.
Collectively, these observations suggest that PKH67 may be used as a specific dye for the labelling
and tracing of SMV1, and perhaps for SSV2. As the viral-host infection experiments were carried
out at 78°C, our study demonstrates the stability of the PKH67 labelling at high temperatures, thus
indicating PKH67 as an option for fluorescent labelling used for tracking organisms above
physiological temperatures.
3.3. Viral uptake by human endothelial cells and assessment of cellular safety
Uptake and intracellular processing of labelled viruses were followed in two human
endothelial cells of different origin. The results in Figure 3A & B show that SMV1 favourably
interact with hCMEC/D3 cells in a concentration-dependent manner, where the proportion of cells
containing viruses also increases with increasing SMV1 concentration at 24 h post incubation. In
contrast to SMV1, SSV2 poorly recognizes hCMEC/D3 cells even at high viral concentrations.
However, both viruses were able to interact with HUVECs (Figure 3C & D), thus suggesting the
presence of additional antigens/receptors on HUVECs that can recognize SSV2 rather than a
mechanism modulated by viral shapes. Whether viral adhesion to and recognition by both cell types
is mediated through the viral tail regions remain to be elucidated.
Live-cell microscopy, however, confirmed viral internalization in both cell types (Figure 4).
These studies further revealed predominant localization of viruses to the peri-nuclear region, and
particularly to lysosomal compartments, and therefore consistent with classical clathrin-mediated
81
endocytic processes [16,20]. The latter process is further supported by viral co-localization to early
endosomes, at early time points (not shown). Due to strong fluorescent signal at the peri-nuclear
region, we also investigated viral trafficking to endoplasmic reticulum (ER) and the Golgi complex.
In contrast to the lysosomal compartment, Manders’ fluorescence overlap coefficient remains low
in both ER and Golgi complex and regardless of the virus type (Figure 4). Therefore, it is unlikely
that these viruses translocate to these organelles effectively, and therefore, internalization routes
such as caveolae-independent and lipid raft/caveolae-mediated endocytosis [21,22], presumably,
operate poorly in relation to archaeal viral uptake, at least in these cells.
Neither SMV1, nor SSV2 induced any deleterious cytotoxic effects. This notion is
confirmed from the apparent lack of detrimental effects on cell morphology, LDH release and
mitochondrial functionality (Figure 5) irrespective of virus load, type and the endothelial cell type.
More specifically, the viral challenge showed no significant changes on basal respiration or
mitochondrial proton leak or maximum respiratory rates (MRR) in either cell lines (Figure 5). This
supports that the physiological coupling state of mitochondrial respiration remains unaltered [17,18]
even following cell exposure to high concentrations of either SMV1 or SSV2. This suggestion is
further verified by calculations of the coupling efficiency of oxidative phosphorylation (OXPHOS),
revealing no significant changes (Table 1).
3.4. Vector development
Anaerobic archaea has been detected in the human oral, colonic and vaginal microbial flora
demonstrating their ability to colonize the human host [5,23-25]. However, with respect to the
safety issue, archaeal viruses have never been reported to integrate into human or any other
eukaryotic genomes, and archaeal viral sequences have never been detected in the sequences of
eukaryotic genomes. This can be explained by the fact that archaeal viral integration needs the
82
presence of an attachment site and a specific integrase enzyme, which cannot be expressed from the
native archaeal viral genome in the eukaryotic cellular environment. Thus, as with plant viruses and
bacteriophages [26], archaeal viruses are less likely to trigger negative downstream effects in
mammals due to an inability to proliferate.
In terms of engineering viruses for development as nanovectors, two major techniques are
implored. These include genetic engineering and surface functionalization (3). Although no
attempts have been made to genetically modify SSV2 or SMV1, a closely related virus to SSV2 has
been shown to be amenable to insertion of genetic material without loss of function (15). Genome
comparison between SMV1 and three related genomes, all retrieved from hot spring metagenomes,
revealed that some genes are not present in all. For example, SMV3 has a much larger genome than
SMV1, this suggests that large DNA fragments could be inserted into the genome of SMV1 without
loss of function. Moreover, several archaeal viruses have been made into genetic model systems,
proving the possibility of genetic engineering of archaeal viruses in general (16). In addition to
genetic engineering, chemoselective chemistry is a powerful tool for incorporating functionalities
onto virus scaffolds. Often naturally occurring amino acid residues, suitable for functionalization
with an appropriate group, are present on the viral particle surface. The crystal structures of SMV1
and SSV2 have not been resolved, however, with the use of iterative threading assembly refinement
we have predicted the 3-dimensional structure of the capsid proteins from the respective amino acid
sequences (Supplementary Figure S1). The results show the presence of several surface functional
groups on the viral capsids such as Lys, Asp, and Glu for chemoselective modification, thus making
it ideal for future surface engineering initiatives.
Another important limitation to using viruses as nanovectors is their physical stability.
For many of the contemplated applications, the improvement of the physical stability of viral
nanoparticles may be critical to adequately meet the demanding physicochemical conditions they
83
may encounter during production and/or storage (17). The extremophilic nature of archaeal viruses
gives them an inherent stability under a wide range of conditions (7, 18). To demonstrate this, we
froze a SMV1 virus stock for 24 h at -20°C without the use of any cryoprotectants. This treatment
did not incur any significant morphological changes to the virus, and viral infectivity (determined
by a plaque assay) was comparable to the native untreated virus (Supplementary Figure S2).
4. Conclusions and perspective
Our results provide the first insights into the uptake by and behaviour of archaeal viruses in human
endothelial cells in vitro. In particular, SSV2 differentiates between HUVECs and hCMEC/D3
cells, thus opening a path for selective cell targeting. The specific recognition and uptake of SMV1
by hCMEC/D3 cells is interesting and may provide an alternative approach for in vivo targeting of
brain cerebral capillary endothelial cells. However, further studies are needed to determine whether
viral interaction with mammalian cells is mediated through their tail regions as well as
identification of cell surface receptors that participate in viral binding and internalization steps. We
also need to learn about the interaction between archaeal viruses and key elements of the innate
immunity including the complement system, blood leukocytes, tissue macrophages and dendritic
cells [6]. Such studies will not only determine the fate of archaeal viruses on intravenous injection,
but also provide insights into viral safety as well as their targeting ability outside the
reticuloendothelial system [6]. Nevertheless, the unique shapes, extremophilic, and acid-resistant
nature of archaeal viruses, together with their unprecedented intracellular safety (at least with
respect to the two tested endothelial cells), could provide new opportunities for development of
sensors for long-term monitoring of biological processes. This may include development of viralbased pH sensors for time-resolved pH measurements in endo-lysosomal compartments, for
instance through surface coupling of pH-sensitive fluorophores to the functional groups of the viral
84
capsid. These approaches may aid our understanding of acidification processes that may control
antigen processing and presentation in relation to dendritic cell maturation [31]. Here, archaeal
viruses may further ferry antigens to such cells through controlled surface manipulation strategies.
However, future attempts should also include and introduce manipulative approaches pertaining the
viral core for encapsulation of biologics including their controlled delivery and release at target
sites. Taken together, our findings suggest that archaeal viruses may hold great promise for future
bioengineering initiatives and development of safe multifunctional vectors and theranostics.
Acknowledgments
The work was supported by The Danish Council for Independent Research /Natural Sciences (grant
number DFF-4181-00274B). SMM acknowledges financial support from Danish Agency for
Science, Technology and Innovation, reference 09-065736 (Det Strategiske Forskningsråd).
hCMEC/D3 cells were obtained under license from Prof. P-P. Couraud, University Paris 05, CNRS,
Institute Cochin, INSEMB, UMR 8104, F-75270 Paris, France.
Competing interests
The authors declare no competing financial interest.
85
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Figure 1. Physical characteristics of viruses before and after labelling with PKH67. (A) SMV1, (B) SMV1-PKH67,
(C) SSV2, (D) SSV2-PKH67. Left panel: Transmission electron micrographs of viruses; Middle: Typical size
distribution profile and concentration of viral preparations determined by Nanoparticle Tracking Analysis; Right panel:
2D plots of relative light scattering intensity of viruses versus the estimate of their size.
88
Figure 2. Host cell imaging of S. islandicus REY 15A infected with SMV1. Cells were stained with DAPI and SMV1
particles were labeled with PKH67 (Panel A). The co-localization of DAPI stained cells (blue, left panel) with virus
particles (green, middle panel) appears as light purple (right panel). Bar = 90 µm. The insets in (A) represent magnified
(2x) views of the selected regions. The Pearson's and Mander's overlap coefficients are represented as the average of 6
individual images of infected cells at 5 mp.i. and 30 mp.i. at an MOI of 10 (Panel B). S. acidocaldarius is not a known
host to any archaeal viruses and was used as a control, and no co-localization was observed when DAPI stained S.
acidocaldarius cells were infected with SMV1-PKH67 at 30 mp.i at an MOI of 10 (not shown). The infectivity was
measured by a plaque assay before and after labelling, and presented as PFU/mL (Panel C).
89
Figure 3. Cellular uptake of PKH67-labelled viruses by hCMEC/D3 cells (A, B) and HUVECs (C, D). Left panel:
Average fluorescence intensity of cells after exposure to increasing concentration of viruses for 24 h (A, C); Right
panel: Quantification of positive cells bearing PKH67-labelled viruses by FACS (B, D). In the all experiments,
corresponding cells were incubated with different concentration of either SMV1-PKH67 or SSV2-PKH67 for 24 h.
90
Figure 4. Intracellular trafficking of PKH67-labelled viruses in hCMEC/D3 cells and HUVECs. Intracellular
trafficking (live-cell tracking) of nanoparticles in different organelles after 24 h post-treatment with SMV1-PKH67 (A
& D) and SSV2-PKH67 (B & E) viruses. Panels (A & B) represent hCMEC/D3 cells and panels (D & E) correspond to
HUVECs cells. Differential interference contrast (DIC) images were taken simultaneously to show morphological
changes and fluorescence positioning. Scale bar = 20 µm. Panels (C) and (F) represent Manders’ overlap coefficient
after image analysis showing the extent of viral-derived fluorescence overlap with the four intracellular compartments
in hCMEC/D3 cells and HUVECs, respectively.
91
Figure 5. Investigation of cytotoxic effect of viruses in hCMEC/D3 cells and HUVECs. Panel (A) shows the extent of
lactate dehydrogenase (LDH) release from cells at 24 h post virus treatment. Panels (B–D) shows the effect of viral
concentration on cell respiratory states at 24 h post incubation (n= 6). Respiration (indicated as the rate of oxygen
consumption) was evaluated at basal (the physiological coupling state controlled by cellular energy demand) (panel B),
Leak (where respiration is independent of ADP phosphorylation and mainly occurs due to proton leak from
mitochondrial intermembrane space) (panel C), and maximum respiratory rate (MRR) states (panel D). Blank =
untreated cells.
92
Supporting information
Figure S1. Prediction of 3-dimensional structure of the capsid proteins of SMV1 and SSV2 from
their respective amino acid sequences by iterative threading assembly refinement (I-TASSER). A
virus capsid is composed of coat proteins, which assemble into the virion structure to protect the
viral DNA. The virus capsid can be composed of one or more coat proteins. Previous studies have
identified the coat proteins present in the archaeal viruses SMV1 and SSV2. The former virus has
two identified coat proteins; ORF 122 and ORF 153 [1,2], and SSV2 also has two identified coat
proteins; VP1 (ORF 88b) and VP3 (ORF 92) [3].
I-TASSER (http://zhanglab.ccmb.med.umich.edu/I-TASSER) was used to identify the availability
of functional groups on the surface of SMV1 (panel A) and SSV2 (panel B). The I-TASSER server
is an integrated platform for automated protein structure and function prediction based on the
sequence-to-structure-to-function paradigm. Starting from an amino acid sequence, I-TASSER first
generates three-dimensional atomic models from multiple threading alignments and iterative
structural assembly simulations. The function of the protein is then inferred by structurally
matching the 3D models with other known proteins [4,5].
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viruses of Archaea: a new way of doing viral business. J Virol 2015; doi:10.1128/JVI.00612-15.
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immune systems of Sulfolobus. Mol Microbiol 2014;91:900-17.
3. Stedman KM, She Q, Phan H, Arnold HP, Holz I, Garrett RA, et al. Relationships between
fuselloviruses infecting the extremely thermophilic archaeon Sulfolobus: SSV1 and SSV2. Res
Microbiol 2003;154:295-302.
4. Roy A, Kucukural A, Zhang Y. I-TASSER: a unified platform for automated protein structure
and function prediction. Nature Protocols 2010;5:725-738.
5. Zhang Y. I-TASSER server for protein 3D structure prediction. BMC Bioinformatics 2008; 9:40.
93
A
94
B
95
Figure S2. Transmission electron micrographs of SMV1 before and after one freeze (-20°C, 24 h)thawing cycle in the absence of cryoprotectants. The viral preparation was suspended in Tris-HCl
buffer, pH 6, then divided into two Eppendorf tubes. The control was stored at 4°C for 24 h. The
frozen sample was stored at -20°C for 24 h without any use of cryoprotectants. The virus sample
was thawed atroom temperature. The virus titer was then calculated by plaque assay and virus
morphology was observed by TEM. Freeze thawing did not induce any apparent morphological
changes. Nanoparticle Tracking Analysis showed a narrow size distribution for SMV1 with a mean
sphere equivalent hydrodynamic diameter of 121 ± 1 nm (n = 3), and 129 ± 6 nm (n = 3) before and
after freeze-thawing cycle. There was no alteration in host infectivity (PFU/mL = 3 x 1010 before
and after freeze-thaw cycle).
96
III SMV1, an extremely stable thermophilic platform for nanoparticle trafficking in the mammalian GI tract Kristine B. Uldahl, Seth T. Walk, Stephen C. Olshefsky, Xu Peng, and Mark Young
Manuscript submitted to PlosOne
97
98
SMV1, an extremely stable thermophilic virus platform for nanoparticle
trafficking in the mammalian GI tract
Kristine B. Uldahl1, Seth T. Walk2, Stephen C. Olshefsky2, Xu Peng1, Mark J. Young3*
1
Danish Archaea Centre and Department of biology, University of Copenhagen, Copenhagen,
Denmark
2
Department of Microbiology and Immunology, Montana State University, Bozeman, Montana,
USA
3
Thermal Biology Institute and Department of Plant Sciences and Plant Pathology, Montana State
University, Bozeman, Montana, USA.
* Corresponding author
E-mail: [email protected] (MY)
99
Abstract
Symbiotic microorganisms of the mammalian GI tract (gut microbiome) play important
roles in maintaining human health, but can also promote the onset and/or progression of certain
diseases. Given the large inter-individuality of microbiome composition, specific GI microbes are
important novel targets for personalized medicine. A major challenge to manipulating microbes in
the GI tract is the safe delivery of effector molecules (i.e. drugs). Nano-vectors, or small molecular
complexes, have been optimized as drug delivery vectors. However, survival during passage
through the extremely acidic, proteolytic, and reductive environment of the alimentary tract poses a
significant challenge. To overcome this challenge, we investigated the acido-thermophilic virus,
Sulfolobus monocaudavirus 1 as a candidate therapeutic nano-vector for the distal mammalian GI
tract microbiome. We investigated the anatomical distribution, vector stability, and immunogenicity
of this virus following oral ingestion in mice and compared these traits to the more classically used
Inovirus vector M13KE. We found that the extremophilic nature of SMV1 particles were highly
stable under both simulated GI tract conditions (in vitro) and in mice (in vivo). We also found that
SMV1 virus could not be detected in tissues outside the GI tract, it elicited a nearly undetectable
inflammatory response, and challenged mice showed no observable signs of pain or distress.
Finally, we used human intestinal organoids (HIOs) to show that labeled SMV1 virus did not
invade or otherwise perturb the human GI tract epithelium. To the best of our knowledge, this is the
first study to evaluate an archaeal virus as a potential therapeutic nanoparticle delivery system. Our
results provide some assurance of the safety of archaeal viruses as mammalian nano-vectors and
provide strong evidence that these viruses have intrinsic and favorable in vivo characteristics for
bioengineering applications, such as drug and vaccine delivery.
100
Introduction
Sophisticated and individualized drugs are needed to mitigate complex human diseases. For
example, the importance of the gut microbiome for human health and during certain diseases has
been established, but the diversity of the microbial community in the human GI tract is vast
(averaging 1013-1014 cells with an estimated 5000 species [1]) and is largely individual-specific.
Therefore, the gut microbiome is a potential new target for personalized drug therapy [2, 3].
Optimal personalized therapies may take the form of drug- or vaccine-delivery vectors on the same
size scale as microorganisms (nano-vectors), which could be non-invasively administered.
However, development of oral nano-vectors for the human gut remains challenging. In particular,
nano-vectors must be stable in diverse chemical environments remaining intact when they reach the
desired body site, which for orally administered vectors, could mean survival in extremely acidic
and proteolytic environments (e.g. pepsins, proteases, and other degradative enzymes of the GI
tract). The intestinal mucus layer is another obstacle; many nanoparticles are trapped in the mucus
and are rapidly removed [4]. Also, nano-vectors used specifically for microbiome therapy should
not cross the epithelial gut barrier and enter into systemic circulation so as to prevent therapeutic
complications due to immunogenicity and off-target effects [5]. Further, when developing
therapeutic nano-vectors, the ease of production, up-scaling, and chemical/biochemical
modification are all important factors to consider.
Viruses are naturally occurring nanoparticle platforms and despite their disease-causing
ability in some contexts, they have many desirable attributes for novel nanobiotechnological
applications. In the field of biomedicine, plant and bacterial viruses (bacteriophages) are favored as
they are non-pathogenic to animals, which limits safety concerns [6]. Cowpea mosaic virus and
other virus-based protein cage architectures have been engineered for viral delivery of tissue- or
cell-specific fluorescent reporters and drugs [7, 8]. In some instances, chemical modifications were
101
added to the virus to increase evasion of the host immune system, for increasing circulation time,
and for increasing tissue distribution [9]. Often, there is a significant disconnect between the in vitro
development of novel viral nano-vectors and their translation into the clinic because promising in
vitro attributes are not maintained in vivo [10]. Thus, evaluating potential nano-vectors in vivo is a
critical step in the development of these novel therapeutics.
To our knowledge, no thermostable archaeal virus (i.e. viruses replicating in the domain
Archaea) has been examined as a potential viral based protein cage nanoparticle vectors in
mammalian systems.
This study addresses the use of Sulfolobus monocaudavirus 1 (SMV1) as a
candidate therapeutic nano-vector for targeting microbial cells of the mammalian gut. SMV1 infects
strains of the hyperthermophilic archaeal genus Sulfolobus, and was originally isolated from an
acidic hot spring (pH 2-3 and 75-80°C) in Yellowstone National Park [11]. The lemon-shaped
SMV1 particles are enveloped and have a diameter of about 120nm and tail fibers protruding from
one end (see Fig 1 for TEM image). Due to their extremophile nature (hyperthermophile/acidresistant), SMV1 particles are promising candidates for surviving passage through a mammalian
gut. As with plant viruses and bacteriophages, SMV1 should not be pathogenic to humans or
animals, cross the gut epithelium, or stimulate a robust host immune response. To our knowledge,
an archaeal virus has never been evaluated in vivo, but the potential for their use as nano-vector
platforms has been proposed [12-14]. We first compared the in vitro stability of SMV1 in simulated
gastric conditions against the more classically used Inovirus vector, M13KE. We then quantified
viral fecal shedding in mice following oral challenge as well as viral infectivity after passage
through the GI tract. In parallel, we examined pro-inflammatory cytokine levels in serum of orallychallenged mice to quantify possible host immune response. Finally, to generate more humanrelevant data, we used human intestinal organoids (HIOs) and fluorescently-labeled SMV1 particles
to determine whether the virus readily crossed the human gut epithelium. Our results provide strong
102
evidence that SMV1 is extremely stable in the mammalian GI tract, it does not stimulate the murine
immune system, and it does not readily cross the human intestinal epithelium.
Materials and methods
SMV1 preparation
SMV1 was propagated in a highly susceptible strain, S. islandicus CRISPR deletion mutant
delta C1C2 [15]. The host culture was grown in Sulfolobus medium supplemented with 0.2%
tryptone, 0.1% yeast extract, 0.2% sucrose and 0.002% uracil (TYS+U medium) [16]. Cultures
were started from -80°C stock; cells were transferred to 50mL TYS+U medium and incubated at
78°C. After 24h of propagation, the 50mL cell culture was transferred to 950mL of pre-heated
(78°C) TYS+U medium. The culture was grown to an OD600 of 0.2-0.3 (typically 24h) at which
time point the host culture was infected with SMV1 suspension. The supernatant containing the
virus particles were collected 48-72 h post infection and concentrated by ultrafiltration using
VivaSpin centrifugal concentrators (Sartorius, Aubagne Cedex, France). Additionally, the virus
concentrate was purified by ultracentrifugation through a 10-40% continuous Iodixanol gradient.
Continuous gradients were prepared by sequentially layering 10, 20, 30 and 40% iodixanol solution
(OptiPrep™, Axis-Shield PoC AS, Oslo, Norway) in 10mM Tris-acetate, pH 6 into 14mL
centrifuge tubes (Beckman Coulter UK Ltd., High Whickham, Bucks, UK), the gradients were
allowed to diffuse overnight at 4 °C in the dark. Ultrafiltrated virus sample was layered over the
continuous gradient (1mL) and centrifuged in a SW-41 rotor (Beckman Coulter) for 6h at
35,000 rpm at 4 °C. The opaque virus band was recovered and an additional ultrafiltration step was
performed; the virus fraction was washed three times with 10mL 10mM Tris-acetate, pH 6 to
remove the Iodixanol from the solution to prevent interference with downstream processes. TEM
103
was used to confirm the presence of virus particles in the recovered fraction. The virus sample was
then stored at 4° until used.
Fluorescent labeling of SMV1 particles
Immediately before labeling, 1µl of PKH67 dye (Sigma, St. Louis, MO) was dissolved in
1mL of Diluent C (Sigma, St. Louis, MO). 2 volumes of diluted PKH67 was added to 1 volume of
virus suspension (250µl) and mixed by pipetting. After 3 min the labeling was stopped by adding 3
volumes of 1% BSA, then incubated for 1 min to allow binding of excess dye. The virus particles
were centrifuged at 17,000 g for 20 min at 4°C, then the supernatant was carefully removed. The
virus particles could be seen as a tiny yellow pellet. The virus particles were re-suspended in 3mL
Tris-acetate, pH 6 buffer and centrifuged at 15.000 RPM for 20 min. The washing step was repeated
twice. After the final wash the virus pellet was re-suspended in 250µl Tris-acetate buffer, pH 6. The
virus titer after labeling was estimated by plaque assay. SMV1 PKH67 particles were stored at 4°C
in the dark until use (stable for at least ½ year).
Determination of SMV1 plaque forming units
The approximate virus titer of unlabeled and fluorescently labeled SMV1 particles was
determined by plaque assays. Serial dilutions (10 µl) of a virus suspension were mixed with a
sample (2mL) of the exponentially growing host culture (2 x 108 cells). The mixtures were
incubated for 30 min at 78°C to allow the adsorption of the virus to the host cells. Immediately
following the addition of 2 mL of TYS+U medium containing 0.25% Gelrite (78°C), the sample
was layered over a premade 0.8% Gelrite plate (78°C). The plates were incubated for 2 days at
78°C. SMV1 plaques appeared as small clear halos easy to enumerate. The final SMV1
concentration averaged between 109–1011 plaque forming units (PFU) per mL.
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Bacteriophage M13KE preparation
Ph.D.-11 phage display peptide library kit was purchased from New England Bioloabs
(Ipswich, MA, USA). The kit contained Escherichia coli (E. coli) host strain, ER2738 and
bacteriophage M13KE. The host strain and phage were propagated according to the detailed
manufacturer protocol. To ensure high virus titer the phage amplification was performed in 2 x
20mL host cultures. The final eluted phage solutions were serially diluted and quantified by plating
on lawns of ER 2738 grown on agar containing 5-bromo-4-chloro-3indolyl-β-D-galactopyranoside
(X-Gal) and isopropyl- β-D-thiogalactopyranoside (IPTG) (LB/IPTG/X-Gal plates), which show
blue plaques after incubation at 37°C overnight. The plaques were then counted estimating the
concentration. The final phage concentration averaged between 1011–1013 PFU/mL.
Stability of SMV1 particles
10µl of a highly concentrated suspension of SMV1 (3 x 107 PFU) were added to 990µl of
the storage medium. Suspensions of ethanol/water and DMSO/water were mixed in proportions 1:1
for a final concentration of 50%. Simulated gastric fluid (SGF) pH 1.6 and simulated intestinal fluid
(SIF) pH 6.5 were prepared according to Klein (2010)[17]. Both solutions were used within 48
hours. The tube contents were mixed by mild vortexing and the tubes were immediately placed in a
37°C incubator simulating the temperature of the human body. As a reference 10 µl of SMV1 was
added to 990µl of Tris-acetate buffer, pH 6. One suspension was stored at 37°C, the other at 4°C
(normal long-term storage temp.). Duplicates of each of the 6 different storage conditions were
stored for 7 days. The suspensions were monitored by plaque assays 1 and 8 hours after incubation
and then once a day for the following 6 days.
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Ethics statement
This study was carried out in strict accordance with the recommendations in the Guide for
the care and use of laboratory animals of the National Institutes of Health. The protocol was
approved by the Institutional Animal Care and Use Committee of Montana State University. All
mice were euthanized by osoflurane inhalation, and all efforts were made to minimize suffering.
Experimental animals
Nine-week-old, male C57BL/6 mice were used for these experiments. Experimental groups
of mice were housed in separate cages. Three mice per bacteriophage M13KE, unlabeled archaeal
virus SMV1, fluorescently-labeled SMV1 (SMV1 PKH97) and Tris buffer controls were prepared
as described above and introduced to each mouse via oral gavage. Each mouse received a 200µl
volume of either viral suspension or buffer. After gavage, mice were observed and fecal pellets
were collected for 5 days. The dose of each virus was as follows: SMV1 = 3.2 x 1011 PFU, M13KE
= 7 x 1011 PFU, and SMV1 PKH67 = 4 x 1010 PFU. Fecal pellets were collected from each mouse at
0, 1, 3, 5, 7, 11, 24, 48, 72, 96, and 120 hours after oral gavage. Immediately after collection, pellets
were stored at -20°C. Mice were euthanized after 5 days and tissue samples (stomach, small
intestine, and colon) were removed for a histopathological analysis. An identical experiment was
conducted for cytokine analysis and 1 mouse from each group was euthanized at 6 and 12 hours
post-gavage. Blood was collected immediately after euthanasia by perfusing the carotid artery with
physiological saline.
Monitoring for signs of disease/distress
Each mouse, in all 3 groups, was examined for lack of movement, ataxia, hunched posture,
ruffled fur, hyperthermia, dehydration, diarrhea, seizure, and death. The mice were examined 5
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times in the first 12 hours after the oral administration then once a day for the following 4 days.
Body weights were recorded on a daily basis.
Extraction of virus particles from fecal pellets following virus
administration
Prior to extraction each individual pellet was weighed for later normalization. The
individual pellets averaged 0.03g and were homogenized in 20mL of SM buffer (50 mM Tris-HCl,
pH 7.5, containing 100 mM NaCl, and 8 mM MgSO4). Particulate matter was removed by
centrifugation at 5000 RPM for 10 min and the supernatant was filtered through 0.32 µm supor
membrane filters (Acrodisc® Syringe Filters, Pall Life Sciences, Corp., East Hills, NY). The virus
present in the filtrate was concentrated by ultrafiltration using 100K MWCO Spin-x centrifugal
concentrators (Corning Incorporated). The final volume was adjusted to 500µl. The viral nucleic
acid (vDNA) was extracted from 200µl concentrate using the PureLink viral RNA/DNA mini kit
(Invitrogen, USA), according to the manufacturer’s protocol. The isolated vDNA was quantified by
qPCR. Additionally, the concentrates of extracted viral particles were tested for infectivity using
plaque assays.
Isolation of vDNA from mouse tissues
Total DNA was extracted from 25 mg brain, liver, spleen (10 mg), and cecal tip tissues,
based on the DNeasy blood and tissue kit (Qiagen, Valencia, CA). We examined tissues from a total
of six mice, groups 1-3, at 6 and 12 hours post virus or buffer administration. All samples were
processed according to the manufacturer's protocol; lysing overnight in 180 µL ATL lysis buffer as
outlined in the DNeasy user manual (“Purification of Total DNA from Animal Tissues” as found in
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DNeasy Blood & Tissue Handbook, Qiagen, Valencia, CA; July 2006). DNA was recovered in a
single elution step with 100 µL AE solution from the kit.
qPCR assays
After isolating vDNA from fecal samples, tissues and serum, qPCR was used to estimate the
approximate amount of SMV1 or M13KE particles in each specific sample. Primer sets specific to
the two viruses were created for the assays. M13KE primers and conditions were previously
published [18]. SMV1 primers were designed using SnapGene software. The primer pair was as
follows:
SMV1F(5’-CCTATTCAACGTATCAAATCCG-3’)
and
SMV1R(5’-
CTTATCGCCTCAACTACTTTATGAAG-3’). The qPCR reaction mixtures (20 µL) consisted of
10 µL of SsoFastTM EvaGreen® Supermix (Bio-Rad), 0.8 µL forward primer (10 µM), 0.8 µL
reverse primer (10 µM), template virus DNA preparation (2 µL) and sterile MilliQ water (6.4 µL).
Amplification was performed with Rotor-Gene Q real-time PCR cycler (Qiagen). SMV1 qPCR
cycling conditions were 95°C (5 min), followed by 30 3-step cycles at 95°C (5 s), 61°C (5 s) and
72°C (20 s), and a final cycle at 72°C (5 min), see Jaye et al. (2003) for M13KE cycling conditions.
Samples from the control mice gavaged with Tris buffer were used as negative controls and all
samples were run in triplicates. Plasmids containing the relevant template DNA were constructed to
establish qPCR standard curves. Each amplicon was cloned into the pCR2.1-TOPO® vector (TOPO
TA-cloning kit, Invitrogen), plated and subsequently, individual clones were picked and grown in
3mL LB medium containing ampicillin. The following day, the plasmids were purified using
PureYeildTM plasmid miniprep (Promega). Plasmids were sequenced to check for potential errors.
Larger batches of correct plasmid DNA were prepared and the concentrations were determined by
spectrophotometry. To generate standard curves, the plasmids were serially diluted. A standard
curve consisted of at least 5 different dilutions ranging between 10 ng/µl – 100 fg/µl, and was
108
included on every run in duplicate. Real-time data capture was performed with Qiagen’s RotorGene Q operating software. The fecal samples with signal above detection limit were screened by
TEM for viral particles (see below).
Cytokine ELISA
Blood was collected from the heart using a syringe, loaded in a serum separator tube and
immediately centrifuged at 10,000 RPM for 5 min. The separated serum was stored at -20°C until
tested for 12 different pro-inflammatory cytokines using a Multi-Analyte ELISArray Kit (Qiagen,
CA, USA) according to manufactures protocols.
Histopathological analysis
After the completion of the experimental time course, three mice were randomly picked for
histopathological analysis, one from each treatment group and anaesthetized using isoflurane. The
stomach, small intestine and colon were collected, fixed in 4% PBS paraformaldehyde and stored at
4°C. After 48 hours the organs were transferred to 70% ethanol. Tissue sections were cut into 5 µm
thick sections, mounted on glass slides, stained with hematoxylin and eosin (H&E) and examined
by a pathologist, screening for apoptotic cells in sections from stomach, small intestine and colon.
SMV1 infection of cultured human intestinal organoids
Human intestinal organoids (HIOs) were obtained from the laboratory of Jason R. Spence,
University of Michigan. Generation, growth and maintenance of HIOs was carried out as previously
described [19]. HIOs were grown to 2.5-3mm in diameter and inoculated with virus suspension
using thin-walled glass capillaries, pulled to fine-point needles. Approximately, 3µl SMV1 PKH67
virus suspension (2 x 109 PFU) or mock (10mM Tris-acetate buffer, pH 6) was injected using
separate needles for each HIO. Immediately after inoculation, HIOs were suspended in fresh
109
intestinal growth medium. Inoculated HIOs were incubated at 37°C and 5% CO2 for 8 hours. At 8
hours post-inoculation, HIOs were washed three times in PBS and fixed with 2.5% gluteraldehyde
in 0.1M sodium phosphate buffer, overnight at 4°C. After fixation, HIOs were washed (PBS) and
embedded in optimal-cutting-temperature (OCT) freezing compound, and frozen at -20°C to make
blocks for sectioning. Sections (10µm) of HIOs were cut using a cryostat and stained for
immunofluorescence analysis. DAPI was used as nuclear counterstain, 10-6 : 1 PhalloidinTetramethylrhodamine B isothiocyanate (TRITC) / PBS was used to identify filamentous actin, and
the alcian blue / PAS stain kit (Newcomer supply) was used to stain epithelial mucins.
Transmission electron microscopy
Virus containing fractions were adsorbed onto carbon-coated copper grids for 5 min and
stained with 2% uranyl acetate. Images were recorded using a Zeiss Leo 912-Omega transmission
electron microscope (operating at 100KV) with a ProScan 2048 X 2048-CCD camera.
Results
SMV1 retains stability and infectivity after exposure to simulated
gastric conditions
SMV1 was selected as a potential nano-vector based on its extremophilic nature. A
concentrated stock of SMV1 particles (10µl containing 109 PFU/mL) were diluted into 990µl of the
storage solution and incubated for 7 days at 37°C. The stability of the particles was monitored by
using plaque assays to estimate any drop in infectivity over time. Overall, the infectivity of SMV1
particles in 1:1 DMSO/H2O and the two buffer control groups (4° and 37°C) remained constant at
107 PFU/mL over the 7 days (Fig. 2). In contrast, the infectivity of SMV1 particles incubated in 1:1
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EtOH/H2O dropped reproducibly after 1 hour and was eliminated by 24 hours. A difference in
SMV1 infectivity was observed (p<0.01) between incubation in SGF (pH 1.6) and SIF (pH 6.5).
SMV1 particles incubated in SIF showed a two-fold drop in infectivity over the 7 days, though a
significant difference was not observed until 120 hours and onwards compared to the Tris 37°C
control, whereas SMV1 particles incubated in SGF showed a three-fold drop in infectivity over the
first 72 hours and no infectivity after this point. We next compared the stability of SMV1 to
bacteriophage M13KE under similar conditions. M13KE phage particles incubated in SGF at 37°C
showed no infectivity at the three tested time points (1, 24, and 72 hours) (S1 Fig.), suggesting that
strong acid is rapidly detrimental to M13KE. However, only a small drop in infectivity was
observed upon incubation in SIF compared to the control (S1 Fig.). In accordance with its
hyperthermophilic nature, we also found that SMV1 infectivity was unaffected by high heat (95°C
for 5 minutes), whereas M13KE was completely inactivated (data not shown). Overall, these
results indicate that SMV1 is much more stable under simulated gastric conditions than M13KE.
Trafficking of SMV1 and M13KE through the murine GI tract
Fecal shedding profiles for SMV1 and M13KE were investigated in mice over a 5-day
period. No indications of pain or distress were observed and mouse weights did not change
significantly in either group (S2 Fig.). Fecal SMV1 shedding showed a reproducible profile (Fig. 3)
with signal above detection limit beginning at 5 hours post-oral gavage. For inter-individual
comparison, viral genome copy number per pellet was standardized to the average pellet weight
(0.03g) over the course of the experiment. The viral shedding peaked at 7 hours and dropped below
the detection limit by the 24 hour time-point, thus a shedding period between 5-12 hours post oral
gavage was observed. The detection was based on the total recovered fraction of viral nucleic acids
in the fecal samples, using sequence-specific primers to detect our viruses. Overall, the same order
of magnitude of SMV1 genomes was detected in fecal samples (1011) as compared to oral gavage
111
dose (1011). To confirm the qPCR results and to test whether the recovered virus fractions were still
infectious, plaque assays were performed. The plaque assays were performed at 5 and 7 hours,
corresponding to the highest virus shedding detected by qPCR (Fig. 4). Passage through the GI tract
appeared to have a significant influence on SMV1 infectivity, as the abundance of PFU per pellet
was low (103) compared to qPCR quantification of SMV1 genome equivalents per pellet (1010).
However, TEM images of recovered SMV1 particles from fecal pellets revealed intact morphology,
which supports the notion that SMV1 particles are excreted in a stable form (see S3 Fig. for TEM
images). Finally, to test whether modified SMV1 particles behave similar to wild type SMV1
particles, we performed an identical GI tract passage experiment with fluorescently labeled SMV1
particles (SMV1 PKH67). SMV1 PKH67 particles had an identical shedding profile compared to
the unmodified virus (Fig. 3). In contrast, SMV1 was much more stable after passage through the
murine GI tract compared to M13KE. For example, the starting administered dose of M13KE
particles was at the same order of magnitude as the dose of SMV1 (1011), but an order of magnitude
lower M13KE particles (1010) were detected in fecal samples compared to SMV1 (1011)(Fig.3).
Also, detection was limited to a narrower time span between 5-7 hours after administration and no
infectivity was observed from plaque assays. Further, we were not able to capture TEM images of
excreted M13KE particles.
From the data above, we estimated the cumulative amount of SMV1 particles shedded as a
percentage of the original dose (Fig. 3). This estimation was based on the average concentration of
SMV1 particles detected in the pellets collected during the peak shedding period and a published
stool frequency estimate of 7 pellets/hour for C57BL/6 mice [20-22]. Based on this approach, a
high percentage (∼80%) of the original SMV1 particles traveled through the entire murine GI tract
and was recovered in the fecal samples. In contrast, less than 10 % of the original administered
112
M13KE particles were excreted in a stable form (Fig. 3)(see S1 Table for calculations). Overall
these results indicate that SMV1 can transverse the mouse GI track more successfully than M13KE.
Detection of vDNA in off-target tissues/organs
To determine whether SMV1 and M13KE particles trafficked to tissues outside the murine
GI tract, we collected and analyzed different tissues for the presence of vDNA by qPCR. Brain,
liver, spleen, cecum, cecal tip, and serum were analyzed from one animal in each treatment group 6
and 12 hours post-oral gavage. Cardiac perfusion was used prior to tissue dissection to exclude
virus that may have been present in the bloodstream. No SMV1 or M13KE vDNA was detected in
the sham controls and no SMV1 vDNA was detected in any tissue/organ. M13KE vDNA was
detected in the cecal tip at both 6 and 12 hours and the signal (Ct) was comparable between these
time-points (Table 1).
Inflammatory cytokine profiling
Serum levels of 12 different pro-inflammatory cytokines were quantified to determine
whether oral administration of SMV1 triggered a pro-inflammatory host immune response (Table
2). Serum was collected from an individual in each of the treatment groups above at 6 and 12 hours
post oral gavage. All cytokines were below the limit of detection, with the exception of a slight
increase in IL1β in the mouse that received M13KE and a slight increase in IFN-γ in the mouse that
received SMV1. Both of these signals were above the limit of detection at 6 hours post-gavage, but
back below the limit of detection at 12 hours post-gavage.
SMV1 particle interaction with human intestinal epithelial cells
To test whether SMV1 particles were taken up by human epithelial cells, we inoculated
concentrated viruses (2 x 109 PFU) into the lumen of human intestinal organoids (HIOs). These
113
structures were derived from embryonic stem cells into 3-dimensional spheres, representing a single
layer of columnar epithelium. HIOs contain the five major cell types found in the human small
intestine, including absorptive enterocytes, goblet cells, enteroendocrine cells, Paneth cells, and
self-renewing stem cells. This monolayer encloses an internal lumen and is surrounded by a layer of
mesenchymal cells on the basal surface [23]. HIOs were inoculated with either the concentrated
SMV1 suspension or Tris buffer, incubated for 8h, and prepared for confocal microscopy (see
methods section). Very few green fluorescent SMV1 PKH67 particles were observed in the HIO
sections, but some particles were visibly associated with the mucus layer on the surface of the HIO
epithelium (Fig.5). No particles were observed inside or between epithelial cells, suggesting that
uptake, if present, was a rare event.
Histopathological analysis
A single mouse from each treatment group was examined for histologic signs of
inflammation. As seen in F, tissue from the stomach, small intestine, and colon were similar in
virus-treated and control mice. No differences were noted between these tissues by a trained
pathologist.
Discussion
The main objective of this study was to evaluate the stability of the acido-thermophilic
archaeal virus SMV1 in the GI tract as a first step towards developing this virus as nanoplatform for
biomedical applications in the gut. SMV1 was chosen due to its extreme thermal stability, lack of
potential hosts in animals and its ease of large-scale production. SMV1 extremophilic nature
suggests it can survive the diverse and harsh conditions present in the GI tract of humans and
animals. In order to evaluate SMV1 against a commonly used viral nanoplatform, we compared it to
114
the E. coli bacteriophage, M13KE, which is the most commonly used phage for in vivo phage
display experiments [24] and as an addressable nanoprotein cage architecture [25]. SMV1 showed
no decline in infectivity when incubated in 50% DMSO. Stability in DMSO is advantageous, as this
chemical is often used as a solvent in bioconjugation protocols [26, 27]. Storage in 50% EtOH
proved to be the most damaging condition for SMV1 particles with infectivity eliminated after 24
hours. Ethanol is a useful solvent when working with silica, which is used for surface
functionalization of nanoparticles with nanosilica in applied nanotechnology [28]. However, if a
two-fold drop in infectivity can be tolerated, short experiments (1-8 hours) appear feasible. We
have shown that SMV1 particles lose infectivity under certain conditions but the virions may
maintain structural integrity providing a non-infectious protein cage, which could prove useful in
certain types of applications.
Viral stability in simulated GI conditions is valuable for prediction of in vivo behavior of
drug formulations [29] and potential viral nanoplatform-based vectors, like phages [24]. Two types
of media were used to evaluate the stability of SMV1 particles; SIF (simulation of small intestinal
conditions in the fasted state), and SGF (simulation of stomach conditions in the fasted state). The
stability of SMV1 measured in our experiments (only a two-fold drop in infectivity in SIF over the
7 days and three-fold drop in infectivity in SGF over 72 hours) provides confidence that this virus
can easily survive passage through the GI tract. For example, the stomach emptying of mice and
humans as quantified by 27Al nuclear magnetic resonance was 50 and 30 minutes [30], respectively,
and the total GI transit time in mice and humans has been approximated to 6-10 hours and 2-4 days
respectively [31-33]. Thus, our in vitro results suggest that SMV1 is likely to remain stable and
infectious in the GI tract of both of these mammals. Finally, SMV1 was much more stable than the
115
commonly used M13KE in both SIF and SGF, suggesting that SMV1 would be more capable of
oral delivery and passage through the GI tract..
The in vivo stability of SMV1 and M13KE was determined by quantifying both viral
genomic DNA by qPCR and viral infectivity by plaque assays. For qPCR we reasoned that viral
DNA would be protected in vivo when packaged inside intact viral capsids thus providing a good
indicator for intact viral particles. The fecal clearance interval of orally administered virus particles
was comparable to previous noted GI transit times (see above) with peak viral shedding 7h post
sample gavage. The viral shedding profiles of SMV1 and M13KE particles occurred in comparable
time intervals indicating similar GI transit times between the two viruses. However, the recovery
and PCR detection in fecal samples of significantly more SMV1 DNA as compared to M13KE
DNA provides strong evidence that more particles of this virus remained intact through the GI tract.
Approximately 80% of the administered SMV1 was recovered in fecal samples. Our finding that
<10% of M13KE particles were present in excreted fecal pellets, suggests that this phage is easily
degraded and may not be sufficiently stable to carry effector functions to distal GI tract targets. In
addition to the stability of the unlabeled SMV1 virus, we also found that a fluorescently-modified
variant had similar passage dynamics, suggesting that chemical modifications of the viral envelope
has minimal or no negative effect on the stability, providing encouragement that future cellular
targeting ligands can be incorporated onto the surface of the SMV1 particles. Taken together with
the observed stability in 50% DMSO, the modification potential of SMV1 particles without loss of
stability appears promising.
We found no evidence that labeled SMV1 were taken up by epithelial cells of human
intestinal organoids. These results suggest that SMV1 particles reaching the human intestines are
116
likely to travel lumenally with minimal epithelial cell interaction. Further support for this
hypothesis is that no SMV1 DNA was detected outside the GI tract, whereas M13KE DNA was
detected in two cecal tip samples.
An important consideration when using a proteinaceous virus particle is the potential for a
robust host immune response. As no archaeal viruses are known to be pathogenic to mammals, we
expected to observe a minimal, if any, inflammatory response at the administered dose. We were
encouraged to find that nearly all of the 12 proinflammatory cytokines were below our limit of
detection, indicating that mice were highly tolerant of both SMV1 and M13KE. One limitation of
our study is the low numbers of mice considered. However, multiple lines of evidence supported
little or no murine response to viral challenge, making additional animal experiments difficult to
justify. The further lack of histopathology (Fig. 6), signs of overt pain or distress, or changes in
weight all suggest that ingestion of SMV1 and M13KE was well tolerated by these mammalian
hosts. Future studies are needed to develop SMV1 particles with the ability to attach to GI tract
targets, such as absorptive enterocytes lining the GI mucosal surface.
Conclusions
To the best of our knowledge, this is the first study to evaluate an archaeal virus as a
potential therapeutic nanoparticle delivery system. We demonstrated that SMV1 particles are
exceptionally stable under simulated GI tract conditions. We also showed that SMV1 particles are
stable in the GI tract of mice, elicited a nearly undetectable inflammatory response, and were well
tolerated.
Although, several viral-based nano-platforms are in development, SMV1 particles
provide many favorable in vivo characteristics for bioengineering applications, such as drug and
vaccine delivery.
117
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119
Figures
Fig. 1. Characteristics of the archaeal virus SMV1 and the bacteriophage M13KE.
Fig. 2. Survival of SMV1 particles. The infectivity of SMV1 after incubation at 37°C in; 50%
DMSO, 50% EtOH, SIF pH 6.5, and SGF pH 1.6. Incubation in Tris-acetate buffer pH 6.0 was
monitored as a reference at two temperatures; 4°C and 37°C. The number of plaque forming units
per milliliter (PFU/mL) was determined by plaque assay and is shown as a function of time in
hours.
120
Fig. 3. Duration of SMV1 (A) and M13 (B) shedding as a function of the viral copy number
determined by qPCR. Stool samples were collected at; 0, 1, 3, 5, 7, 11, 24, 48, 72, 96 and 120
hours, all were assayed by qPCR (24h are depicted). The peak period was defined by signal above
the detection limit (1E-03 copies/pellet). The viral copy number pr. pellet was standardized to a
pellet of 0.03g. The Tris control group showed no positive signal at any time-point. The red dotted
line indicates the cumulative amount of virus shedded over the peak period as a percentage of the
total administered concentration. The cumulative amount is based on the average viral conc. in the
pellets from 1 mouse collected over the peak period with a stool frequency of 7 pellets/hr (the
cumulative curves were comparable for all mice in the group).
121
Fig. 4. Duration of SMV1 shedding as a function of the plaque forming units pr. pellet. Stool
samples were collected at; 0, 1, 3, 5, 7, 11, 24, 48, 72, 96 and 120 hours, all were assayed by plaque
assays. Duration was defined by the time interval in which the samples provided plaques. The PFU
/ pellet was standardized to a pellet of 0.03g. The Tris control group was negative at all tested timepoints.
Fig. 5. SMV1 particle interaction with human intestinal epithelial cells. The uptake of SMV1
particles in human intestinal organoids was evaluated by confocal microscopy. SMV1 were
fluorescently labeled with the green dye PKH67, the cell nuclei were stained blue with DAPI, the
actin filaments were red, and the mucus dye appear black.
122
Fig. 6. Histological examination of tissues from mice challenged with SMV1 and M13KE.
Histopathological observation of tissues obtained from stomach, small intestine and colon,
demonstrating no treatment-related microscopic effects. A. Tris-acetate control; B. Bacteriophage
M13; C. Archaeal virus SMV1.
123
Table. 1. Detection of SMV1 and M13KE DNA in tissues of mice inoculated orally with virus.
Tissue
Perfused
SMV1
0/2
0/2
0/2
0/2
0/2
0/2
Brain
Liver
Spleen
Cecum
Cecal tip
Serum
Perfused
M13
0/2
0/2
0/2
0/2
2/2
0/2
Control
0/2
0/2
0/2
0/2
0/2
0/2
Control
0/2
0/2
0/2
0/2
0/2
0/2
Table 2. Inflammatory cytokine profiling in serum of mice inoculated orally with virus
Conc. In 100µl mouse serum (the ELISA was run on a 10E-1 dilution)
Cytokine
IL1 α
IL1 β
IL2
IL4
IL6
IL10
IL12
IL17A
IFN-γ
TNF-α
G-CSF
GM-CSF
0,07
0,089
0,054
0,049
0,108
0,061
0,086
0,052
0,06
0,058
0,064
0,119
6h Tris
-0,08
-0,01
-0,06
0,02
-0,22
0,13
-0,21
0,28
0,03
0,02
0,07
-0,12
6h SM1
-0,02
0
-0,12
-0,07
-0,37
0,12
-0,2
0,05
0,87
-0,02
0,09
0,07
6h M13
0,06
0,6
0,06
0,05
-0,24
0,14
-0,06
0,31
0,05
0,02
-0,04
0,12
12h Tris
-0,01
-0,2
0
0,04
-0,29
0,13
-0,19
0,12
0,16
0,18
-0,02
-0,04
12h SMV1
0,05
-0,04
0,13
0,12
-0,2
0,2
-0,14
0,26
0,13
0,2
0,11
0,07
12h M13
0,04
0
0,04
0,06
-0,29
0,16
-0,15
0,2
-0,11
0,21
0,01
0,11
1,70
0,29
1,82
Neg
Pos
1,65
1,05
1,83
1,87
1,13
0,83
1,41
2,45
>2.5
a
The serum levels of 12 pro-inflammatory cytokines measured in experimental animals at timepoints: 6h and 12h under identical conditions.
a
Absorbance values >2.5 are not within the linear range of the assay. Lower limit of detection;
values less than two times the negative control should not be interpreted. Absorbance read at
450nm.
124
Supporting information
S1 Fig. Survival of M13KE particles. The infectivity of M13 after incubation at 37°C in; SIF pH
6.5 and SGF pH 1.6. Incubation in TBS buffer pH 7.5 was monitored as a control. The number of
plaque forming units per milliliter (PFU/mL) was determined by plaque assay and is shown as a
function of time in hours.
S2 Fig. Observations of mice body weights. Body weights were measured daily for 5 days. At the
0h time-point a one-time administration of either 200µl VLP solution or control Tris-acetate
solution was carried out.
125
S3 Fig. TEM images of recovered SMV1 particles from the fecal samples show intact
morphology.
S1 Table. Estimation of virus titer. Estimated SMV1 titer in original inoculum and those detected
in fecal pellets. Estimates based on qPCR and PFU assays are presented.
126
IV Review: Emerging nanoplatforms based on extremophilic archaeal viruses Kristine Buch Uldahl and Xu Peng
Manuscript in preparation formatted as 1400 word review article
127
128
Review: Emerging nanoplatforms based on extremophilic archaeal viruses Kristine B. Uldahl and Xu Peng Viruses are everywhere. Viruses populate virtually every known ecosystem on the planet, including the extreme cold, acidic, and saline environments. They can even be found cruising in the upper atmosphere (1). As a group, archaeal viruses constitute the smallest group of known viruses. These viruses are often morphologically and genetically unique with exceptional life-­‐cycle traits (2). To date, all archaeal viruses characterized, infect extremophilic Archaea (Table 1 and Table 2). Their extreme habitats are often characterized by environmental fluctuations and temporal variations (3). Just as their hosts, archaeal viruses have evolved to thrive under these extreme environmental conditions, making them extremely stable nanoparticles. Initially the unique morphotypes of archaeal viruses stole the spotlight, but in recent years interest has arisen to their application potential, as their extremophilic nature offers unique opportunities in terms of nanoengineering. Although traditionally recognized for their roles as infectious agents, viruses have been engineered over the past decade as highly promising nanoplatforms for the targeted delivery and treatment of human diseases (4). Their attractive features include their nanometer range size, a natural propensity to self-­‐assemble into distinct shapes, their exceptional stability, robustness, biocompatibility and biodegradability. Moreover, viral nanoparticles (VNPs) can be chemically or genetically engineered to exhibit specific functionalities for therapeutic effect (5). Plantbased viruses and bacteriophages are typically considered safer nanoplatforms than mammalian viruses because they cannot proliferate in humans and hence are less likely to trigger adverse effects (6). Archaeal viruses fit this profile as well however their potential remains largely untapped. The design and implementation of viruses in nanomedicine is still in its early stages, with only very few examples in preclinical trials. One of the major challenges is that until virus assemblies are tested in vivo, their cellular uptake, intracellular trafficking and accumulation, together with pharmacokinetics, toxicity and immunogenicity cannot be predicted (4). Thus, 129
in order to develop a virus into a therapeutic nanoplatform in depth knowledge of the virus both in vitro and in vivo is essential. Until recently, no studies had evaluated an archaeal virus in vivo making their proposed potential solely hypothetical. A new study by Uldahl et al. (2015) found that the archaeal virus, SMV1, was highly stable in both simulated conditions of the human GI tract (in vitro) and when passaged orally in mice (in vivo). Further, SMV1 virus particles could not be detected in tissues outside the murine GI tract; it elicited a nearly undetectable murine inflammatory response; challenged mice showed no observable signs of pain or distress; and it did not invade or become intimately associated with the human epithelium when tested in human intestinal organoids. The stability of SMV1 was compared to the traditionally used bacteriophage vector, M13KE. SMV1 outperformed this state-­‐of-­‐the-­‐art vector as measured by in vitro and in vivo survival (replication). In addition, SMV1 was just as immunotolerant in mice as M13KE. These results provide strong evidence that SMV1 in particular and archaeal viruses in general have intrinsically favorable in vivo characteristics for bioengineering applications. Most progress with nanovector delivery of therapeutics has been made with eukaryotic viruses such as adenovirus, which have an intrinsic tropism for human cells and are naturally evolved to overcome mammalian cellular barriers (7). Bacteriophages have no intrinsic tropism for mammalian cell receptors but have been modified to display tissue-­‐specific ligands on the coat protein without disruption of their virus structure thus providing them with target specificity (8). However, this approach encounters problems such as stability issues of the virus-­‐ligand complex in vivo, steric hindrance of the virus uptake by large virus-­‐
ligand complexes, toxicity issues, and limitations to scale up the vector production (9, 10). Interestingly, another study by Uldahl et al. (2015) showed that the two archaeal viruses, SMV1 and SSV2, was taken up by two endothelial cell lines of human origin without any prior modifications. SMV1 showed a differential uptake profile with a high uptake by the brain microvascular endothelial cell line hCMEC/D3, a cell line that can serve as a useful in vitro model of the blood brain barrier (11). This preliminary study demonstrates that the native tropism of SMV1 provide intrinsic specificity for endothelial brain cells that potentially could be explored for the delivery of therapeutics to the brain. 130
In terms of engineering viruses for development as therapeutic nanovectors, two techniques are mainly used, genetic engineering and surface functionalization (6). Although no attempts have been made to genetically modify an archaeal virus for nanomedical purposes, several archaeal viruses have been made into genetic model systems, proving the possibility of genetic engineering of archaeal viruses in general (14). Also, the archaeal virus SSV1 has been shown to be amenable to insertion of genetic material without loss of function (15). Metagenomic analysis suggests that SMV1 may be amenable for genetic insertions as well. Genome comparison of 4 closely related SMV strains revealed that some genes are not present in all. For example, SMV3 has a much larger genome than SMV1, suggesting that large DNA fragments could be inserted into the genome of SMV1 without loss of function (16). In addition to genetic engineering or by itself, chemoselective chemistry is a powerful tool for incorporating functionalities onto virus scaffolds. Often naturally occurring amino acid residues, suitable for functionalization with an appropriate group, are present on the viral particle surface. For example, chemical conjugation of SIRV2 nanoparticles can be controlled selectively and spatially by the type of chemistry used; amine-­‐specific labelling occurs exclusively at the minor capsid protein, selectively labelling the tail regions of the SIRV2 particles (5). Figure 1 illustrates different modification strategies using SMV1 as a potential nanoplatform. Overall, archaeal viruses offer distinct advantages when compared to the plant viruses and bacteriophages. When designing efficient carrier systems, physical properties such as size, morphology, and surface charge, of the nanocarrier exterior have direct influence on cellular uptake and intracellular trafficking. As a group, archaeal viruses exhibit the most diverse morphotypes, providing more alternatives to choose between than any other virus group. Their unique shapes include; lemon-­‐shape, bottle-­‐shape, and droplet-­‐shape (Fig. 2) (12). Moreover, lack of stability has been reported to be problematic for synthetic nanoparticles. Studies have tested the inherent stability of both the halophilic virus His1 and the acido-­‐
thermophilic viruses SIRV2 and SMV1 showing all three viruses to remain infectious under a range of extremely harsh conditions (5, 13, PlosOne), suggesting these nanoparticles are perfectly suited for harsh industrial processes. 131
Other archaeal viruses that could prove interesting to investigate for distinct stability features would be viruses isolated from deep-­‐sea hydrothermal vents. These viruses have adapted to extreme pressure and heat, which could possibly provide nanoplatforms suitable for autoclaving processes. It would also be interesting to evaluate archaeal viruses with exceptional shapes such as bottle-­‐, droplet-­‐, and coil-­‐shape, as it has been reported that morphology or shape is of higher importance than size during initial cell uptake and internalization (4, 17). In total, 83 archaeal viruses have been characterized and are available for evaluation as potential nanoplatforms. As shown in table 1 + 2, all known archaeal viruses are extremophiles belonging to one of two groups. The hyperthermophilic viruses represent the largest group (47, Table 1) and have been isolated from hot acidic springs, solfataric mudholes, deep-­‐sea hydrothermal vents, and anaerobic sludge digesters, whereas halophilic viruses (36, Table 2) originate from hypersaline environments such as The Dead Sea. Although the known archaeal viruses reveal an exceptional degree of diversity with regard to both morphotypes and genomes, this might still be an underrepresentation as they have only been isolated from a limited number of host species (60 out of 83 archaeal viruses infect one of four archaeal genera) and geographic locations. Notably, no viruses infecting psychrophilic or mesophilic Archaea have been characterized. Therefore screening for viruses in other phylogenetic taxa of the archaeal domain could provide even larger diversity in terms of shapes and sizes for future applications. Over the last decade viral nanoparticles have taken giant strides towards the clinic and their development is accelerating. To date, focus has been put on plant viruses and bacteriophages. Here, we advocate the inclusion of archaeal viruses as therapeutic nanovectors based on the promising pilot studies of the archaeal virus SMV1. The exceptional stability of archaeal viruses against various harsh treatments (temperature, pH, pressure, solvents etc.), their unique morphologies and the observed outstanding behaviour of SMV1 in mammalian systems make them a highly promising group of candidates for the future development of novel nanoplatforms. 132
References 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. Sharoni S, Trainic M, Schatz D, Lehahn Y, Flores MJ, Bidle KD, et al. Infection of phytoplankton by aerosolized marine viruses. Proceedings of the National Academy of Sciences of the United States of America. 2015;112(21):6643-­‐7. Prangishvili D, Forterre P, Garrett RA. Viruses of the Archaea: a unifying view. Nat Rev Microbiol. 2006;4(11):837-­‐48. Anderson RE, Sogin ML, Baross JA. Evolutionary strategies of viruses, bacteria and archaea in hydrothermal vent ecosystems revealed through metagenomics. PloS one. 2014;9(10):e109696. van Kan-­‐Davelaar HE, van Hest JCM, Cornelissen JJLM, Koay MST. Using viruses as nanomedicines. Brit J Pharmacol. 2014;171(17):4001-­‐9. Steinmetz NF, Bize A, Findlay KC, Lomonossoff GP, Manchester M, Evans DJ, et al. Site-­‐specific and Spatially Controlled Addressability of a New Viral Nanobuilding Block: Sulfolobus islandicus Rod-­‐
shaped Virus 2. Adv Funct Mater. 2008;18(21):3478-­‐86. Yildiz I, Shukla S, Steinmetz NF. Applications of viral nanoparticles in medicine. Current opinion in biotechnology. 2011;22(6):901-­‐8. Verma IM, Weitzman MD. Gene therapy: twenty-­‐first century medicine. Annual review of biochemistry. 2005;74:711-­‐38. Yata T, Lee KY, Dharakul T, Songsivilai S, Bismarck A, Mintz PJ, et al. Hybrid Nanomaterial Complexes for Advanced Phage-­‐guided Gene Delivery. Molecular therapy Nucleic acids. 2014;3:e185. Cardinale D, Carette N, Michon T. Virus scaffolds as enzyme nano-­‐carriers. Trends in biotechnology. 2012;30(7):369-­‐76. Buning H, Ried MU, Perabo L, Gerner FM, Huttner NA, Enssle J, et al. Receptor targeting of adeno-­‐
associated virus vectors. Gene therapy. 2003;10(14):1142-­‐51. Weksler B, Romero IA, Couraud PO. The hCMEC/D3 cell line as a model of the human blood brain barrier. Fluids and barriers of the CNS. 2013;10(1):16. Nasir A, Forterre P, Kim KM, Caetano-­‐Anolles G. The distribution and impact of viral lineages in domains of life. Frontiers in microbiology. 2014;5:194. Hong C, Pietila MK, Fu CJ, Schmid MF, Bamford DH, Chiu W. Lemon-­‐shaped halo archaeal virus His1 with uniform tail but variable capsid structure. Proceedings of the National Academy of Sciences of the United States of America. 2015;112(8):2449-­‐54. Wirth JF, Snyder JC, Hochstein RA, Ortmann AC, Willits DA, Douglas T, et al. Development of a genetic system for the archaeal virus Sulfolobus turreted icosahedral virus (STIV). Virology. 2011;415(1):6-­‐11. Stedman KM, Schleper C, Rumpf E, Zillig W. Genetic requirements for the function of the archaeal virus SSV1 in Sulfolobus solfataricus: construction and testing of viral shuttle vectors. Genetics. 1999;152(4):1397-­‐405. Gudbergsdóttir SR, Menzel P, Krogh A, Young M, Peng X. Novel viral genomes identified from six metagenomes reveal wide distribution of archaeal viruses and high viral diversity in terrestrial hot springs. Environmental microbiology. 2015;in press. Truong NP, Whittaker MR, Mak CW, Davis TP. The importance of nanoparticle shape in cancer drug delivery. Expert opinion on drug delivery. 2015;12(1):129-­‐42. 133
3
2
1
Fig 1. Virus engineering scheme of potential surface modifications of SMV1 with different ligands. (1) Genetic insertion of a foreign bioactive peptide sequence into the capsid. (2) The foreign peptide can serve as a handle for bioconjugation reactions introducing ligands such as fluorophores and affinity tags. (3) Wild-­‐type virus capsid proteins can also be chemically modified using bioconjugation protocols. In general care is needed to avoid the disruption of virus properties. Fig 2. Electron micrographs of (A) ABV (Haring et al. 2005); (B) SSV2; (C) SNDV (Arnold et al.
2000); (D) SMV1. Scale bars 50 nm in C; 100 nm in A and B; 200 nm in D.
134
Table 1 ⏐ Characteristics of hyperthermophilic archaeal virus Morphology Virus
Host
Origin
Genome
References
Ampullaviridae
Bottle-shaped
ABV
Acidianus
Hot acidic spring
(93°C, pH 1.5)
Italy
Ln
23.9
(Peng et al., 2007)
Bicaudaviridae
Spindle-shaped
ATV
Acidianus
Hot acidic spring
(93°C, pH 1.5)
Italy
C
62.7
(Prangishvili et al.,
2006)
Clavaviridae
bacilliform
APBV1
Aeropyrum
Coastal hot spring
Japan
C
5.2
(Mochizuki et al.,
2010)
Globulaviridae
Spherical
PSV
Pyrobaculum
Hot spring (85°C, pH 6)
Yellowstone, USA
Ln
28.3
(Haring et al.,
2004)
TTSV1
Thermoproteus
Hot spring
Indonesia
Ln
28.3
(Ahn et al., 2006)
APOV1
Aeropyrum
Isolated from culture
strain
C
13.8
(Mochizuki et al.,
2011)
SNDV
Sulfolobus
Solfataric fields
New Zealand
C
20.0
(Arnold et al.,
2000a)
ASV1
Acidianus
Hot acidic spring
(88.3°C, pH 3.5)
Iceland
C
24.2
(Redder et al.,
2009)
PAV1
Pyrococcus
Hydrothermal vents
Fiji Basin
C
18.0
(Geslin et al., 2007)
SMF1
Sulfolobus
Hot acidic spring
(65°C, pH 3.6)
Azufres, Mexico
C
14.8
(ServinGarciduenas et al.,
2013a)
SSV1
Sulfolobus
Hot acidic spring
(80°C, pH 2.5-3)
Beppu, Japan
C
15.5
(Palm et al., 1991)
SSV2
Sulfolobus
Hot acidic spring
Iceland
C
14.8
(Stedman et al.,
2003)
SSV4
Sulfolobus
Hot acidic spring
Iceland
C
15.1
(Peng, 2008)
SSV5
Sulfolobus
Hot acidic spring
Iceland
C
15.3
(Redder et al.,
2009)
SSV6
Sulfolobus
Hot acidic spring
Iceland
C
15.6
(Redder et al.,
2009)
SSV7
Sulfolobus
Hot acidic spring
C
(Redder et al.,
Guttaviridae
Droplet-shaped
Fuselloviridae
Spindle-shaped
135
Hot acidic spring
(75°C, pH 4)
Kamchatka, Russia
C
17.4
(Wiedenheft et al.,
2004)
SSVRH
Sulfolobus
Hot acidic spring
(75°C, pH 4)
Kamchatka, Russia
C
16.4
(Wiedenheft et al.,
2004)
TPV1
Thermococcus
Hydrothermal vents
East Pacific Rise
C
21.5
(Gorlas et al., 2012)
AFV1
Acidianus
Hot acidic spring
(85°C, pH 1.5-2)
Yellowstone, USA
Ln
21.1
(Bettstetter et al.,
2003)
Acidianus
Hot acidic spring
(93°C, pH 2)
Italy
Ln
31.8
(Haring et al.,
2005)
Acidianus
Hot acidic spring
(93°C, pH 1.5)
Italy
Ln
40.5
(Vestergaard et al.,
2008a)
AFV6
Acidianus
Hot acidic spring
(93°C, pH 1.5)
Italy
Ln
39.6
(Vestergaard et al.,
2008a)
AFV7
Acidianus
Hot acidic spring
(93°C, pH 1.5)
Italy
Ln
36.9
(Vestergaard et al.,
2008a)
AFV8
Acidianus
Hot acidic spring
(93°C, pH 1.5)
Pozzuoli, Italy
Ln
38.2
(Vestergaard et al.,
2008a)
AFV9
Acidianus
Hot acidic lake (82°C,
pH 4)
Kamchatka,Russia
Ln
41.2
(Bize et al., 2008)
SIFV
Sulfolobus
Solfataric fields
Iceland
Ln
40.9
(Arnold et al.,
2000b)
(δ)
AFV3
(β)
α-shape
(β)
TTV1
(α)
Thermoproteus
Mud-hole (93°C, pH 6)
Iceland
Ln
15.9
(Janekovic et al.,
1983)
TTV2
Thermoproteus
Mud-hole (93°C, pH 6)
Iceland
Ln
15.9
(Janekovic et al.,
1983)
(β)
Thermoproteus
Mud-hole (93°C, pH 6)
Iceland
Ln
27
(Janekovic et al.,
1983)
SMV1
Sulfolobus
Hot acidic spring
(85°C, pH 2)
Yellowstone, USA
ARV1
Acidianus
Hot acidic spring
(β)
TTV3
Monocaudavirus
Spindle-shaped
2009)
Sulfolobus
AFV2
β-, γ-, δ-shape
17.6
SSVK1
(γ)
Lipothrixviridae
Filamentous
Iceland
136
C
48.8
Ln
(Erdmann and
Garrett, 2012)
(Vestergaard et al.,
Rudiviridae
Rod-shaped
Siphoviridae
Head-tail
Spiraviridae
Coil-shaped
Unclassified
Spherical
Unclassified
Spindle-shaped
24.7
2005)
SIRV1
Sulfolobus
Solfataric fields
Iceland
Ln
32.3
(Prangishvili et al.,
1999)
SIRV2
Sulfolobus
Solfataric fields
Iceland
Ln
35.5
(Prangishvili et al.,
1999)
SRV
Stygiolobus
Hot acidic spring
(93°C, pH 2)
Azores, Portugal
Ln
28.1
(Vestergaard et al.,
2008b)
SMR1
Sulfolobus
Hot acidic spring
(65°C, pH 3.6)
Azufres, Mexico
Ln
27.4
(ServinGarciduenas et al.,
2013b)
ΨM2
Methanobacterium
Anaerobic sludge
digester (60°C)
Switzerland
Ln
26.11
(Meile et al., 1989)
ΨM100
Methanothermobacter
Methanothermobacter
wolfeii culture
Ln
28.8
(Luo et al., 2001)
ACV
Aeropyrum
Hot spring (98°C)
Japan
C
24.9
(Mochizuki et al.,
2012)
STIV1
Sulfolobus
Hot acidic spring
(72-92°C, pH 3-4)
Yellowstone, USA
Ln
17.7
(Rice et al., 2004)
STIV2
Sulfolobus
Hot acidic spring
(88.3°C, pH 3.5)
Iceland
C
17.6
(Happonen et al.,
2010)
ATSV
Acidianus
Hot acidic spring
(80°C, pH 2)
Yellowstone, USA
C
71
(Hochstein et al.,
2015)
HAV1
Hot acidic spring
Yellowstone, USA
Ln
22.7
(Garrett et al.,
2010)
HAV2
Hot spring (85°C, pH 6)
Yellowstone, USA
C
17.7
(Garrett et al.,
2010)
Turriviridae
Icosahedral
Unclassified
Spindle-shaped
(93°C, pH 1.5)
Italy
APSV1
Aeropyrum
Isolated from culture
strain
C
38.1
(Mochizuki et al.,
2011)
STSV1
Sulfolobus
Hot acidic springs and
mudholes
Tengchong, China
C
75.3
(Xiang et al., 2005)
STSV2
Sulfolobus
Hot acidic spring
(88.3°C, pH 3.5)
Iceland
C
17.6
(Happonen et al.,
2010)
137
Table 2 ⏐ Characteristics of halophilic archaeal virus Morphology
Virus
Host
Salterprovirus
Spindle-shaped
HIS1
Haloarcula
Hypersaline lake
Australia
Ln
14.5
HF1
Halorubrum
Solar saltern
Australia
Ln
75.9
(Tang et al., 2004)
HF2
Halorubrum
Solar saltern
Australia
Ln
77.7
(Tang et al., 2002)
HGTV1
Halogranum
Solar saltern
Thailand
C
143.9
(Atanasova et al.,
2012)
HRTV4
Halorubrum
Solar saltern
Italy
Ln
35.7
(Atanasova et al.,
2012)
HRTV5
Halorubrum
Solar saltern
Italy
Ln
76.1
(Atanasova et al.,
2012)
HRTV7
Halorubrum
Solar saltern
Italy
Ln
69.0
(Atanasova et al.,
2012)
HRTV8
Halorubrum
Solar saltern
Thailand
Ln
74.5
(Atanasova et al.,
2012)
HSTV2
Haloarcula
Solar saltern
Israel
Ln
68.2
(Atanasova et al.,
2012)
φCh1
Natrialba
Hypersaline lake
Kenya
Ln
58.5
(Klein et al., 2002)
ΦH
Halobacterium
Hbt. salinarum
culture
Ln
59.0
(Schnabel et al., 1982)
HRPV1
Halorubrum
Solar saltern
Italy
C
7.1
HRPV2
Halorubrum
Solar saltern
Thailand
C
10.7
(Atanasova et al.,
2012)
HRPV3
Halorubrum
The Dead Sea
Israel
C
8.8
(Atanasova et al.,
2012)
HRPV6
Halorubrum
Solar saltern
Thailand
C
8.6
(Atanasova et al.,
2012)
HIS2
Haloarcula
Hypersaline lake
Australia
Ln
16.1
(Bath et al., 2006)
HRPV2
Halorubrum
Solar saltern
Thailand
C
10.7
(Atanasova et al.,
2012)
HSTV1
Haloarcula
Solar saltern
Italy
Ln
32.2
(Atanasova et al.,
2012)
Myoviridae
Head-tail
Pleolipoviridae
Pleomorphic
Podoviridae
Head-tail
Origin
138
Genome
Reference
(Bath et al., 2006)
(Pietila et al., 2009)
Siphoviridae
Head-tail
Sphaerolipoviridae
Tailless
Unclassified
Pleomorphic
BJ1
Halorubrum
Salt lake
Mongolia
Ln
42.3
HRPV3
Halorubrum
The Dead Sea
Israel
C
8.8
(Atanasova et al.,
2012)
HRPV6
Halorubrum
Solar saltern
Thailand
C
8.6
(Atanasova et al.,
2012)
HCTV1
Haloarcula
Solar saltern
Italy
Ln
103.3
(Atanasova et al.,
2012)
HCTV2
Haloarcula
Solar saltern
Thailand
Ln
54.3
(Atanasova et al.,
2012)
HCTV5
Haloarcula
Solar saltern
Thailand
Ln
102.1
(Atanasova et al.,
2012)
Hh1
Halobacterium
Anchovy sauce
Philippines
NT
32.7
(Pauling, 1982)
Hh2
Halobacterium
Anchovy sauce
Philippines
NT
29.4
(Pauling, 1982)
HHTV1
Haloarcula
Solar saltern
Italy
Ln
49.1
(Atanasova et al.,
2012)
HHTV2
Haloarcula
Solar saltern
Thailand
Ln
52.6
(Atanasova et al.,
2012)
HVTV1
Haloarcula
Solar saltern
Thailand
Ln
101.7
(Atanasova et al.,
2012)
ΦN
Halobacterium
Hbt. Halobium
culture
Ln
56.0
(Vogelsang-Wenke
and Oesterhelt, 1988)
HHIV2
Haloarcula
Solar saltern
Italy
Ln
30.6
(Jaakkola et al., 2012)
PH1
Haloarcula
Hypersaline lake
Australia
Ln
28.1
(Porter et al., 2013)
SH1
Haloarcula
Hypersaline lake
Australia
Ln
30.9
(Porter et al., 2005)
SNJ1
Natrimena
Natrimena sp.
culture
C
16.3
(Mei et al., 2007)
HHPV1
Haloarcula
Solar saltern
Italy
C
8.1
(Roine et al., 2010)
HGPV1
Halorubrum
Solar saltern
Spain
C
9.7
(Atanasova et al.,
2012)
139
(Pagaling et al., 2007)
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