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Home , Picornavirus

On the entry and entrapment of

Jacqueline Staring

PS_JS_def.indd 1 08-10-18 08:46 ISBN 978-94-92679-60-4 NUR 100

Printing and lay-out by: Proefschriftenprinten.nl – The Netherlands With financial support from the Netherlands Cancer Institute

© J. Staring, 2018 All rights are reserved. No part of this book may be reproduced, distributed, or transmitted in any form or by any means, without prior written permission of the author.

PS_JS_def.indd 2 08-10-18 08:46 On the entry and entrapment of picornaviruses

Over het binnendringen en insluiten van picornavirussen

(met een samenvatting in het Nederlands)

Proefschrift

ter verkrijging van de graad van doctor aan de Universiteit Utrecht op gezag van de rector magnificus, prof. dr. H.R.B.M. Kummeling, ingevolge het besluit van het college voor promoties in het openbaar te verdedigen op dinsdag 6 november 2018 des middags te 16.15 uur

door

Jacqueline Staring

geboren op 8 december 1984 te Leiden

PS_JS_def.indd 3 08-10-18 08:46 Promotor: Prof. dr T.R. Brummelkamp

PS_JS_def.indd 4 08-10-18 08:46 Table of Contents

Chapter 1 Introduction I: Picornaviruses 7

Chapter 2 Introduction II: Genetic approaches to study 27 host- interactions

Chapter 3 PLA2G16, a switch between entry and clearance of 43 picornaviridae

Chapter 4 D68 receptor requirements unveiled by 77 haploid genetics

Chapter 5 KREMEN1 is a host entry receptor for a major group 95 of

Chapter 6 General Discussion I: Viral endosomal escape & 125 detection at a glance

Chapter 7 General Discussion II: Concluding remarks 151

Addendum 161

Summary 163

Nederlandse samenvatting 165

Curriculum vitae 167

List of publications 168

Acknowledgements 169

PS_JS_def.indd 5 08-10-18 08:46 Roll the dice

if you’re going to try, go all the way. otherwise, don’t even start.

PS_JS_def.indd 6 08-10-18 08:46 1

Introduction I: Picornaviruses

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Poliovirus and the history of molecular virology

The family consists of ipv over more than 94 species1 infecting 1 organisms ranging from , rodents, and cattle, to humans. Most members of the picornavirus family have been extensively studied: specifically, has been, and still is, an archetype that led the way for the development of the field of molecular virology. The disease caused by poliovirus (PV), poliomyelitis, is also known as infantile paralysis and records seemingly depicting the disease have been found dating back as far as ancient Egyptian times2,3. Poliomyelitis is a devastating disease that occurs when poliovirus spreads from the gastro-intestinal tract to the central nervous system of the patient. Most patients suffer from mild symptoms such as fatigue, fever, vomiting, and headaches 4. However, 1-2% of the patients developed more serious illness with symptoms ranging from overall muscle weakness, paralyzed limbs, difficulty breathing and swallowing and in some cases death 5. Although the disease was feared under the general population, real scientific interest in the virus causing poliomyelitis did not develop until the beginning of the 20th century, when the epidemics became more prevalent and severe 6. In 1908 Erwin Popper and Karl Landsteiner were the first to discover that poliomyelitis was a non-bacterial infectious substance, by injecting two monkeys with filtered spinal cord fluid of a 9-year-old patient. Both monkeys developed poliomyelitis type symptoms, proving for the first time that is was a disease transmitted by a virus7 . Previous studies with other model organisms (i.e. rabbits and mice) had failed to demonstrate transmission from infected patient material to animals, implying that only humans and monkeys (in an experimental setting) can serve as a host8. The availability of a suitable model organism to study the virus led to a quick surge in the knowledge on the pathogenicity of PV. Discoveries such as entry routes of PV into the host (through the mucosa of the nose to the central nervous system 9), the identification of the major site of replication (in the gastro- intestinal tract; hence the name enterovirus 10), and, finally, the way the virus spread from patient to patient (through the oral-fecal route 10). After two failed vaccine trails in the 1930s by John Kolmer, Maurice Brodie and William Park (first in monkeys 11,12, later in children 12), and the development of a respirator nicknamed the “iron lung” in 1928 13,14, the next major breakthrough came from Albert Sabin and Peter Olitsky, who were for the first time able to culture PV in embryonic nervous tissues in vitro 15, and later also in cell monolayers 16, enabling de novo synthesis of virions in extracts. Even though vaccine development

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was hampered for almost two decades by the failed vaccine trails, the second largest outbreak in the US in 1952 with an incidence of 35 patients per 100.000 people 6,17 reignited attempts to generate a safe poliovirus vaccine, eventually leading to the first (inactivated poliovirus) vaccine in 1953 by Jonas Salk18 . Studies to find a suitable PV vaccine catapulted the molecular virology field forward. Poliovirus had evolved into a model organism that together with Tobacco Mosaic Virus (TMV), would serve as a valuable research tool for virology research and the coming of age of the field of molecular biology. Soon after the publication of Salk, and later the publication on the life attenuated vaccine of Sabin 19, the focus shifted away from vaccine development towards a need for understanding of both the virus and host. One major topic of interest was the identification of the attachment-, or entry receptor required for PV to infect cells. A first indication that such a receptor existed came from studies from Evans and coworkers, who demonstrated that cells became more susceptible for PV infection when cultured for some time in vitro, whereas cells freshly isolated from tissues did not support infection 20. This suggested that there was a cellular host factor required for infection present on the cell membrane of in vitro cultured cells, which was not present of the freshly isolated tissues. After being able to generate new infectious TMV by transfecting the genomic RNA of TMV into Tobacco plants 21, and the successful transfection of (another picornavirus species) 22 and poliovirus 23, similar experiments with poliovirus RNA in non-permissive cells demonstrated that resistant cells are capable of supporting infection when the entry barrier was circumvented24. This discovery, together with the finding that some tissues support virus propagation better than others 25, started hinting towards an essential cellular attachment/entry factor that could be responsible for the observed tissue tropism during infection 24. The final identification of the PV receptor did not occur until the 1980s after the cloning 26 and sequencing 27,28 of infectious cDNA of the poliovirus , when Mendelsohn et al. decided to transfect a cDNA library from HeLa cells (which support PV infection) in non-permissive mouse L-cells 29. The L-cell clones that now permitted infection were isolated lead to the eventual cloning and sequencing of the poliovirus receptor (PVR) CD155 30. Around the same time, both the X-ray crystal structures of -14 (HRV-14) 31 and poliovirus 32 virions were solved, leading to a further understanding of receptor binding.

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E xpansion of the picornavirus family

For a long period of time, the Picornavirus family was subdivided in 4 genera; 1 Enterovirus, Rhinovirus, , and , based on their sedimentation, stability, and buoyancy 31. However, with the introduction of sequencing techniques, many new picornavirus species were identified and classification based on sequence similarity became the standard. This did not only lead to the renaming, deletion, and addition of genera, serotypes, and species, but also lead to a huge increase in known picornaviruses. The International Committee on Taxonomy of (ICTV) is still updating the picornavirus family tree on a regular basis 33. By now, the Picornaviridae consists of 5 genera known to infect humans (Enterovirus, Hepatovirus, , , and Cardiovirus), and 35 other genera infecting other of which the Aphthovirus genus is studied most.

Genomic organization of picornaviruses

The increase in sequenced picornaviruses species did not only assist in , it did also boost the knowledge of the replication and organization of their genomes. All picornaviruses possess a single, small, positive stranded RNA (pico-RNA) genome of about 7500 nucleotides, with a 5’ non-translated region (NTR), an open reading frame encoding for a single polyprotein, a poly-A tail, and a 3’-NTR 34. This uncapped RNA genome is transported from the viral over the cellular membrane to the cytosol to initiate infection. Once there, it gets directly translated via an internal ribosomal entry site (IRES) mediated translation 35 into a single polyprotein 28, which is subsequently auto-cleaved by viral to form the viral required for genome replication and the subsequent formation of viral progeny. The genome organization of picornaviruses is strikingly similar. Most viruses have a VPg peptide linked to their 5’ NTR of the RNA genome 36,37 followed by a transcript encoding for a polyprotein which is sectioned in precursor regions P1, P2, and P3. These regions consists of a leader protein (L) protein (for cardio-, and ) 38,39, capsid proteins (VP0) VP4, VP2, VP3, VP1 (P1 region); non-structural proteins 2A, 2B (viporin), and replication factor 2C (P2 region); replication factor 3A, replication and translation factor 3B (VPg), protease 3C, and finally the RNA polymerase 3D (P3 region) 40–45. All of which are flanked by the 5’ and 3’ NTRs, of which both sequence as well as the secondary structures play

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an important role during translation initiation and genome replication 46–53 (Figure 1). Of the non-structural proteins, the L proteins, 2A, and 2B are least conserved among picornavirus species 34.

F igure 1. Schematic organization of the picornavirus genome. All picornavirus genomes are approximately 7500 bp long, consisting of a 5’ untranslated region followed by the structural proteins (P1), non-structural proteins (P2 and P3) and the 3’ NTR.

Although genetic drift and recombination between picornaviruses frequently occur, most viruses do not tolerate major deletions or insertions in their genome. This was, for instance, demonstrated with a transposon-based insertion screen in an infectious PV cDNA clone a small (15 nucleotide) marker 54. This screen identified only a few sites in the PV genome in which the sequence is stably inserted while still yielding infectious virus particles. However, markers genes like dsRed or GFP cloned into these site were recombined out within a couple of replication rounds of the mutant viruses55. Similar results were obtained with (CV) B3 mutants in which eGFP was introduced in the beginning of the of a CV- B3 infectious cDNA. Here again, eGFP expression was lost after a few rounds of infection, suggesting that the insertion of larger stretches of sequence in the genome are poorly tolerated by these viruses 56. Whereas large insertions and deletions are not commonly observed, picorna­ viruses are well known to undergo substantial genetic drift, which is believed to be

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beneficial for adaptation strategies57 . This drift is mostly attributed to the lack of proofreading of the 3D RNA polymerase and gives rise to a large array of mutant viruses also known as quasispecies 57,58. The high mutation rate enables the virus 1 to very quickly overcome selective pressure 59 and often hampers drug and vaccine development (of which the development of rhinovirus vaccines are the most notorious example 60,61. Another related characteristic of picornaviruses is their ability to recombine large sections of their genome with serotypes within their species62. These horizontal recombination events were already described in the 60s in experimental settings with different PV mutants 63,64, but have also been described for other entero-, and aphthoviruses 65–67 in vivo. Furthermore, chimeric serotypes are often observed during outbreaks of coxsackie A viruses which arise due to co-infections in one patient with multiple serotypes 68,69. Sequence analysis point out that these recombination events often evolve around “hot spots” mostly just before and after the structural P1 region of the genome 66,69,70 . As mentioned before, the homology between picornavirus genomes of the different genera diverge only on a few locations.O ne of which is the region encoding for the 2A protein(s). Poliovirus and other enteroviruses, for instance, possess one copy of 2A, which functions as a cysteine protease, cleaving both the viral polyprotein 71, as well as cellular host factors such as eukaryotic translation initiation factor 4G (eIF4G) 72,73. However, other picornaviruses such as Ljungan virus are known to have two copies of 2A, which structurally resemble either the 2A from the Cardio-, and Aphthovirus genera, or the 2A proteins of the Parechovirus genus 74, suggesting that picornaviruses evolved from an ancestral virus with serval 2A proteins. Furthermore, Aichi virus (Kobuvirus genus), and (Parechovirus genus) possess a 2A protein lacking the characteristic catalytic triad required for the protease activity of the enterovirus 2A’s. It is suggested that these 2A proteins are RNA binding proteins involved in genome replication 75,76. Strikingly, the proposed catalytic residues of the 2A proteins of these viruses (together with avian encephalomyelitis virus (AEV)), show a higher sequence similarity with a mammalian protein family of phospholipases 77. What’s more, one of these phospholipases, PLA2G16, serves as a host factor for many picornaviruses that possess an enterovirus-like protease version of 2A (chapter 3) 78, but is not required for the replication of parechovirus type 1 (unpublished data). This homology between viral and mammalian proteins suggests that although horizontal recombination events between picornaviruses and their host are extremely rare, they do seem to occur.

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B rief introduction into the picornavirus lifecycle

The genetic material encoding for the viral proteins is encapsidated in the an icosahedral virion of approximately 30 nm diameter, encompassing 60 copies of the four viral capsid proteins (VP1/VP2/VP3/VP4), which destabilizes upon specific queues when the virion has reached its site of cell entry79. While picornaviruses in rare occasions incorporate genes from hosts into their genome, they cannot carry all genetic information required for the formation of new progeny. In consequence, picornaviruses rely heavily on multiple host genes during their infectious life-cycle 34. The next paragraph will shortly describe the general aspects of the lifecycle of picornaviruses. More in depth reviews on this topic can be found elsewhere 34,79,80. While all picornaviruses display a very similar lifecycle, they are known to divert in their dependency on attachment-, and entry receptors required for access to the host cell. TheEnterovirus genus serves as a good example: poliovirus requires CD155 (poliovirus receptor, or PVR) 30, the minor group of (RV) depend on the low density lipoprotein receptor (LDLR) 81, where the major group of RVs depend mostly on ICAM1 82, type B use the Coxsackie and Adenovirus Receptor (CAR) 83, and finally enterovirus-A71 (EV-A71) has a dual dependency on P-selectin glycoprotein ligand-1 (PSGL-1) 84 and scavenger receptor class B member 2 (SCARB2) 85. Moreover, there are enterovirus species of which the receptor remains to be determined (as will be discussed in chapter 4 and chapter 5). After attachment, picornaviruses use various endocytic pathways for uptake, and are known to have different requirements for acidification as a cue to initiate the uncoating process. In short, once arrived on their designated location in the endo- lysosomal compartment, most picornavirus virions undergo a conformational shift prompting the extrusion of, and the subsequent imbedding in the host membrane of the N-terminal part of VP1 86. The myristoylated VP4 is also externalized and embedded into the cell membrane and is completely lost from the virion 87–91, leading to the formation of a 135S or A particle 91–95. This interaction of the viral capsid proteins with the cell membrane leads to a direct pore-based connection with the cytoplasm which is suggested to form without the assistance of other host factors 94,96,97. Nevertheless, it remains to be determined whether this pore formation is sufficient for the formation of a 80S particle after the proper uncoating and release of the RNA genome into the cytoplasm (as shall be discussed in chapter 3) 94. The viral genome needs to reach the cytoplasm to initiate translation, which is solely dependent on, and initiated by, the host through IRES mediated translation.

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The first round of replication yields several viral proteins through autodigestion by the 2A and 3C proteases (and by the Leader protein in the case of aphthoviruses) 98,99, including intermediate protein products that have their own distinct 1 function during infection. The 2A and 3C proteases also initiate the shutdown of and translation of cellular products. This is achieved by the digestion of proteins required for transcription such as TATA box binding protein (TBP) 100, p53 101, cyclic AMP-responsive element binding protein (CREB) 102, and octamer- binding protein (Oct-1) 103 by protease 3C and its precursor protein 3CD 104,105. Cap-mediated translation is restricted by the cleavage of proteins such as poly(A)- binding protein (PABP), eukaryotic translation initiation factor 4A (eIF4A) by 3C protease 106 and eIF4G by 2A protease 73,107,108. This allows for the redirection of all cellular resources to the production of viral progeny while also curbing the anti-viral response of the cell. The newly produced non-structural viral proteins also initiate a massive reengagement of membranous structures of the infected cell. These so-called replication vesicles are required the synthesis of new copies of the viral genome 109– 113. Replication is initiated by the generation of a negative stranded RNA molecule that serves as a template, with the uridylylated form of VPg (VPg-pUpU) as a primer, for the formation of multiple new positive stranded RNA genomes. These newly generated genomes interact already during transcription with capsid proteins, in which they are packaged to form new viral particles 114. Finally, these viruses egress from the cell either through cell lysis 115–117, or through non-lytic spreading in autophagosome-like vesicles 118–121 (Figure 2).

Relevance of picornavirus infections

The vaccination programs for the eradication of polio are one of the major success stories of modern medicine. Poliovirus type 2 is officially eradicated for more than a decade122 and poliovirus type 3 has not been detected since 2012 123. Efforts are now directed towards prevention of vaccine derived poliovirus infections (by discontinuing the use of attenuated poliovirus based vaccines) and towards the final eradication of poliovirus type 1, which would make polio the second disease after to be eradicated as a result of human efforts 123. Although these developments are positive, other picornaviruses (mostly other enteroviruses) are still responsible for severe outbreaks. These viruses have proven to cause serious illness in humans that are equivalent to poliovirus infections124–127. Less prevalent

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F igure 2. Life-cycle of picornaviruses. All picornaviruses share a similar infectious life-cycle. First, they bind to the host cell via a receptor (1) followed by internalization of the receptor-virion complex (2). After encountering different uncoating ques (acidification, receptor binding etc.), picornaviruses modify the membrane of the endosomal compartment (3), which is rapidly followed by the release of the genome from the capsid into the cytoplasm (4). Here, the genome is translated by the host (5), after which the polyprotein gets processed into different viral proteins (6). RNA replication on replication vesicles is initiated (7) and new viral progeny is formed (8), which are subsequently released from the host cell through cell lysis or budding (9).

viruses like EV-D68 have been linked to polio-like-illness 128–130, and more widespread viruses such as EV-A71 and CV-A16, can cause symptoms ranging from hand-foot-and-mouth-disease, to and encephalitis, causing fatalities amongst young children124,126,127,131,132. Furthermore, coxsackieviruses have been linked to myocarditis and (although controversial) to the development of type 1 diabetes133. Some non-enterovirus species such as parechoviruses have been linked to myocarditis and pericarditis 134. Interestingly, the viruses that are of largest concern are the ones that may cause the least (human) fatalities. Rhinoviruses (causing the ) and foot-and-mouth-disease virus (FMDV, a disease

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affecting cloven-hoofed animals) are a major economic burden on a global scale, due to, for instance, missed work days 135 and culling of farm animals 136. Indubitably, existing vaccines and new vaccine developments will aid in the 1 prevention of picornavirus infections. However, one of the major hurdles in combating picornavirus infections and virus infections in general is the absence of antiviral curative therapies against which viruses do not develop resistance. This coincides with our poor understanding of how these viruses manage to escape detection of the (cellular) immune system of their host.

Thesis outline

This Thesis will focus on the host factors that play an important role during picornavirus infections, specifically during entry, with the use of genetic screens in human haloid cells. First, in chapter 2, an overview will be given on which host factor screens are most commonly used to study molecular virology, specifically the haploid genetic screens that are used in the experimental chapters of this thesis, and how these screens have increased our knowledge of virus entry factors in the past for other virus families. Chapter 3 will focus on a phospholipid modifying enzyme, PLA2G16, and its role during entry of multiple picornavirus genera. The next two chapters will demonstrate the power of haploid genetics in the discovery in new entry receptors for EV-D68 (chapter 4) and for a large group of serotypes of the Enterovirus type A species (chapter 5). Finally, chapter 6 will provide an overview on how viruses penetrate the endo/lysosomal compartment to gain entry into the cell and which detection mechanisms the host has evolved to counteract it.

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References

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20. Smith, W. M. & Evans, C. A. Growth of neurotropic viruses in extraneural tissues. VII. Cultivation of MM and Western equine encephalomyelitis viruses in cultures of rhesus testicular tissue. J Immunol 72, 353–359 (1954). 1 21. GIERER, A. & SCHRAMM, G. Infectivity of Ribonucleic Acid from Tobacco Mosaic Virus. Nature 177, 702–703 (1956). 22. COLTER, J. S., , H. H. & BROWN, R. A. Infectivity of Ribonucleic Acid from Ehrlich Ascites Tumour Cells Infected with Mengo Encephalitis. Nature 179, 859–860 (1957). 23. Colter, J. S., Bird, H. H., Moyer, A. W. & Brown, R. A. Infectivity of ribonucleic acid isolated from virus-infected tissues. Virology 4, 522–532 (1957). 24. Holland, J. J., Mc, L. L. & Syverton, J. T. The mammalian cell-virus relationship. IV. Infection of naturally insusceptible cells with enterovirus ribonucleic acid. J Exp Med 110, 65–80 (1959). 25. Holland, J. J. Receptor affinities as major determinants of enterovirus tissue tropisms in humans. Virology 15, 312–326 (1961). 26. Racaniello, V. R. & Baltimore, D. Cloned poliovirus complementary DNA is infectious in mammalian cells. Science 214, 916–919 (1981). 27. Nomoto, A. et al. Complete nucleotide sequence of the attenuated poliovirus Sabin 1 strain genome. Proc. Natl. Acad. Sci. U. S. A. 79, 5793–5797 (1982). 28. Kitamura, N. et al. Primary structure, gene organization and polypeptide expression of poliovirus RNA. Nature 291, 547–553 (1981). 29. Mendelsohn, C. et al. Transformation of a human poliovirus receptor gene into mouse cells. Proc Natl Acad Sci U S A 83, 7845–7849 (1986). 30. Mendelsohn, C. L., Wimmer, E. & Racaniello, V. R. Cellular receptor for poliovirus: molecular cloning, nucleotide sequence, and expression of a new member of the immunoglobulin superfamily. Cell 56, 855–865 (1989). 31. Rossmann, M. G. et al. Structure of a human common cold virus and functional relationship to other picornaviruses. Nature 317, 145–153 (1985). 32. Hogle, J. M., Chow, M. & Filman, D. J. The structure of poliovirus.Sci. Am. 256, 42–49 (1987). 33. ICTV Master Species Lists 2017. Arch. Virol. (2018). 34. Lin, J. Y. et al. Viral and host proteins involved in picornavirus life cycle. J. Biomed. Sci. 16, (2009). 35. Pelletier, J. & Sonenberg, N. Internal initiation of translation of eukaryotic mRNA directed by a sequence derived from poliovirus RNA. Nature 334, 320–325 (1988). 36. Lee, Y. F., Nomoto, A., Detjen, B. M. & Wimmer, E. A protein covalently linked to poliovirus genome RNA. Proc. Natl. Acad. Sci. U. S. A. 74, 59–63 (1977). 37. Kitamura, N. et al. The genome-linked protein of picornaviruses. VII. Genetic mapping of poliovirus VPg by protein and RNA sequence studies. Cell 21, 295–302 (1980). 38. Chen, H. H., Kong, W. P. & Roos, R. P. The leader peptide of Theiler’s murine encephalomyelitis virus is a zinc-binding protein. J. Virol. 69, 8076–8 (1995). 39. Devaney, M. A., Vakharia, V. N., Lloyd, R. E., Ehrenfeld, E. & Grubman, M. J. Leader protein of foot and mouth disease virus is required for cleavage of the p220 component of the cap binding protein complex. J Virol. 1988 Nov; 62(11) 4407 9 (1988).

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75. Samuilova, O., Krogerus, C., Pöyry, T. & Hyypiä, T. Specific interaction between human parechovirus nonstructural 2A protein and viral RNA. J. Biol. Chem. 279, 37822–37831 (2004). 76. Sasaki, J. & Taniguchi, K. Aichi virus 2A protein is involved in viral RNA replication. J. Virol. 82, 9765–9769 (2008). 77. Hughes, P. J. & Stanway, G. The 2A proteins of three diverse picornaviruses are related to each other and to the H-rev107 family of proteins involved in the control of cell proliferation. J. Gen. Virol. 81, 201–207 (2000). 78. Staring, J. et al. PLA2G16 represents a switch between entry and clearance of Picornaviridae. Nature 541, 412–416 (2017). 79. Marsh, M. & Helenius, A. Virus entry: Open sesame. Cell 124, 729–740 (2006). 80. Baggen, J., Jan Thibaut, H., Strating, J. R. P. M. & van Kuppeveld, F. J. M. The life cycle of non- polio enteroviruses and how to target it. Nat. Rev. Microbiol. (2018). doi:10.1038/s41579-018- 0005-4 81. Hofer, F. et al. Members of the low density lipoprotein receptor family mediate cell entry of a minor-group common cold virus. Proc. Natl. Acad. Sci. 91, 1839–1842 (1994). 82. Greve, J. M. et al. The major human rhinovirus receptor is ICAM-1.Cell 56, 839–847 (1989). 83. Bergelson, J. M. et al. Isolation of a common receptor for Coxsackie B viruses and adenoviruses 2 and 5. Science 275, 1320–1323 (1997). 84. Nishimura, Y. et al. Human P-selectin glycoprotein ligand-1 is a functional receptor for . Nat. Med. 15, 794–7 (2009). 85. Yamayoshi, S. et al. Scavenger receptor B2 is a cellular receptor for enterovirus 71. Nat. Med. 15, 798–801 (2009). 86. Strauss, M., Levy, H. C., Bostina, M., Filman, D. J. & Hogle, J. M. RNA Transfer from Poliovirus 135S Particles across Membranes Is Mediated by Long Umbilical Connectors. J. Virol. 87, 3903–3914 (2013). 87. Chow, M. et al. Myristylation of picornavirus capsid protein VP4 and its structural significance. Nature 327, 482–486 (1987). 88. Danthi, P., Tosteson, M., Li, Q. -h. & Chow, M. Genome Delivery and Ion Channel Properties Are Altered in VP4 Mutants of Poliovirus. J. Virol. 77, 5266–5274 (2003). 89. Panjwani, A. et al. Capsid Protein VP4 of Human Rhinovirus Induces Membrane Permeability by the Formation of a Size-Selective Multimeric Pore. PLoS Pathog. 10, (2014). 90. Moscufo, N., Yafal, a G., Rogove, a, Hogle, J. & Chow, M. A mutation in VP4 defines a new step in the late stages of cell entry by poliovirus. J. Virol. 67, 5075–5078 (1993). 91. Fricks, C. E. & Hogle, J. M. Cell-induced conformational change in poliovirus: externalization of the amino terminus of VP1 is responsible for liposome binding. J. Virol. 64, 1934–1945 (1990). 92. Crowell, R. L. & Philipson, L. Specific alterations of coxsackievirus B3 eluted from HeLa cells. J. Virol. 8, 509–15 (1971). 93. de Sena, J. & Mandel, B. Studies on the in vitro uncoating of poliovirus II. Characteristics of the membrane-modified particle.Virology 78, 554–566 (1977).

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94. Tuthill, T. J., Bubeck, D., Rowlands, D. J. & Hogle, J. M. Characterization of early steps in the poliovirus infection process: receptor-decorated liposomes induce conversion of the virus to membrane-anchored entry-intermediate particles. J. Virol. 80, 172–180 (2006). 1 95. Ren, J. et al. Picornavirus uncoating intermediate captured in atomic detail. Nat. Commun. 4, (2013). 96. Tosteson, M. T., Wang, H., Naumov, A. & Chow, M. Poliovirus binding to its receptor in lipid bilayers results in particle-specific, temperature-sensitive channels.J. Gen. Virol. 85, 1581– 1589 (2004). 97. Bilek, G. et al. Liposomal Nanocontainers as Models for Viral Infection: Monitoring Viral Genomic RNA Transfer through Lipid Membranes. J. Virol. 85, 8368–8375 (2011). 98. Ypma-Wong, M. F., Filman, D. J., Hogle, J. M. & Semler, B. L. Structural domains of the poliovirus polyprotein are major determinants for proteolytic cleavage at Gln-Gly pairs. J. Biol. Chem. 263, 17846–17856 (1988). 99. Donnelly, M. L. L., Gani, D., Flint, M., Monaghan, S. & Ryan, M. D. The cleavage activities of aphthovirus and cardiovirus 2A proteins. J. Gen. Virol. 78, 13–21 (1997). 100. Clark, M. E., Lieberman, P. M., Berk, A. J. & Dasgupta, A. Direct cleavage of human TATA- binding protein by poliovirus protease 3C in vivo and in vitro. Mol. Cell. Biol. 13, 1232–7 (1993). 101. Weidman, M. K., Yalamanchili, P., Ng, B., Tsai, W. & Dasgupta, A. Poliovirus 3C protease- mediated degradation of transcriptional activator p53 requires a cellular activity. Virology 291, 260–271 (2001). 102. Yalamanchili, P., Datta, U. & Dasgupta, A. Inhibition of host cell transcription by poliovirus: cleavage of transcription factor CREB by poliovirus-encoded protease 3Cpro. J. Virol. 71, 1220–6 (1997). 103. Yalamanchili, P., Weidman, K. & Dasgupta, A. Cleavage of transcriptional activator Oct-1 by poliovirus encoded protease 3C(pro). Virology 239, 176–185 (1997). 104. Sharma, R., Raychaudhuri, S. & Dasgupta, A. Nuclear entry of poliovirus protease-polymerase precursor 3CD: Implications for host cell transcription shut-off.Virology 320, 195–205 (2004). 105. Amineva, S. P., Aminev, A. G., Palmenberg, A. C. & Gern, J. E. Rhinovirus 3C protease precursors 3CD and 3CD??? localize to the nuclei of infected cells. J. Gen. Virol. 85, 2969–2979 (2004). 106. Kuyumcu-Martinez, N. M., Van Eden, M. E., Younan, P. & Lloyd, R. E. Cleavage of Poly(A)- Binding Protein by Poliovirus 3C Protease Inhibits Host Cell Translation: a Novel Mechanism for Host Translation Shutoff.Mol. Cell. Biol. 24, 1779–1790 (2004). 107. Ventoso, I., MacMillan, S. E., Hershey, J. W. b. & Carrasco, L. Poliovirus 2A proteinase cleaves directly the eIF-4G subunit of eIF-4F complex. FEBS Lett. 435, 79–83 (1998). 108. Sommergruber, W. et al. 2A Proteinases of Coxsackie- and Rhinovirus Cleave Peptides Derived from eIF-4? via a Common Recognition Motif. Virology 198, 741–745 (1994). 109. Schlegel, A., Giddings, T. H., Ladinsky, M. S. & Kirkegaard, K. Cellular origin and ultrastruc- ture of membranes induced during poliovirus infection. J. Virol. 70, 6576–88 (1996). 110. Kallman, F., Williams, R. C., Dulbecco, R. & Vogt, M. Fine structure of changes produced in

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cultured cells sampled at specified intervals during a single growth cycle of polio virus.J. Cell Biol. 4, 301–308 (1958). 111. Bienz, K., Egger, D., Rasser, Y. & Bossart, W. Intracellular distribution of poliovirus proteins and the induction of virus-specific cytoplasmic structures.Virology 131, 39–48 (1983). 112. Caliguiri, L. A. & Tamm, I. Membranous structures associated with translation and transcription of poliovirus RNA. Science 166, 885–6 (1969). 113. Limpens, R. W. A. L. et al. The transformation of enterovirus replication structures: A three- dimensional study of single- and double-membrane compartments. MBio 2, (2011). 114. Belov, G. A. et al. Complex Dynamic Development of Poliovirus Membranous Replication Complexes. J. Virol. 86, 302–312 (2012). 115. Sandoval, I. V & Carrasco, L. Poliovirus infection and expression of the poliovirus protein 2B provoke the disassembly of the Golgi complex, the organelle target for the antipoliovirus drug Ro-090179. J Virol 71, 4679–4693 (1997). 116. Nieva, J. L., Agirre, A., Nir, S. & Carrasco, L. Mechanisms of membrane permeabilization by picornavirus 2B viroporin. in FEBS Letters 552, 68–73 (2003). 117. Van Kuppeveld, F. J. M. et al. Coxsackievirus protein 2B modifies endoplasmic reticulum membrane and plasma membrane permeability and facilitates virus release. EMBO J. 16, 3519–3532 (1997). 118. Wilson, T., Papahadjopoulos, D. & Taber, R. Biological properties of poliovirus encapsulated in lipid vesicles: antibody resistance and infectivity in virus-resistant cells. Proc. Natl. Acad. Sci. U. S. A. 74, 3471–5 (1977). 119. Chen, Y. H. et al. Phosphatidylserine vesicles enable efficient en bloc transmission of enteroviruses. Cell 160, 619–630 (2015). 120. Bird, S. W., Maynard, N. D., Covert, M. W. & Kirkegaard, K. Nonlytic viral spread enhanced by autophagy components. Proc. Natl. Acad. Sci. 111, 13081–13086 (2014). 121. Bird, S. W. & Kirkegaard, K. Escape of non-enveloped virus from intact cells. Virology 479– 480, 444–449 (2015). 122. WHO. Polio Eradication & Endgame Midterm Review July 2015. (2015). 123. WHO. Polio Eradication & Endgame Midterm Review July 2015. (2015). 124. Xing, W. et al. Hand, foot, and mouth disease in China, 2008-12: An epidemiological study. Lancet Infect. Dis. 14, 308–318 (2014). 125. Ho, M. et al. An Epidemic of Enterovirus 71 Infection in Taiwan. N. Engl. J. Med. 341, 929–935 (1999). 126. Liu, S. L. et al. Comparative epidemiology and virology of fatal and nonfatal cases of hand, foot and mouth disease in mainland China from 2008 to 2014. Reviews in Medical Virology 25, 115– 128 (2015). 127. Solomon, T. et al. Virology, epidemiology, pathogenesis, and control of enterovirus 71. The Lancet Infectious Diseases 10, 778–790 (2010). 128. Messacar, K. et al. A cluster of acute flaccid paralysis and cranial nerve dysfunction temporally associated with an outbreak of enterovirus D68 in children in Colorado, USA. Lancet 385, 1662–1671 (2015).

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129. Holm-Hansen, C. C., Midgley, S. E. & Fischer, T. K. Global emergence of enterovirus D68: A systematic review. The Lancet Infectious Diseases 16, e64–e75 (2016). 130. Greninger, A. L. et al. A novel outbreak enterovirus D68 strain associated with acute flaccid 1 myelitis cases in the USA (2012-14): A retrospective cohort study. Lancet Infect. Dis. 15, 671– 682 (2015). 131. Shindarov, L. M. et al. Epidemiological, clinical, and pathomorphological characteristics of epidemic poliomyelitis-like disease caused by enterovirus 71. J. Hyg. Epidemiol. Microbiol. Immunol. 23, 284–95 (1979). 132. Ho, M. et al. An Epidemic of Enterovirus 71 Infection in Taiwan. N. Engl. J. Med. 341, 929–935 (1999). 133. Hober, D. & Alidjinou, E. K. Diabetes: Towards a coxsackievirus B-based vaccine to combat T1DM. Nature Reviews Endocrinology 14, 131–132 (2018). 134. McGregor, T., Bu’Lock, F. A., Wiselka, M. & Tang, J. W. Human parechovirus infection as an undiagnosed cause of adult pericarditis. Journal of Infection 75, 596–597 (2017). 135. Fendrick, A. M., Monto, A. S., Nightengale, B. & Sarnes, M. The economic burden of non- influenza-related viral respiratory tract infection in the United States.Arch. Intern. Med. 163, 487–494 (2003). 136. Knight-Jones, T. J. D. & Rushton, J. The economic impacts of foot and mouth disease - What are they, how big are they and where do they occur? Preventive Veterinary Medicine 112, 162– 173 (2013).

PS_JS_def.indd 25 08-10-18 08:46 if you’re going to try, go all the way. this could mean losing girlfriends, wives, relatives, jobs and maybe your mind.

PS_JS_def.indd 26 08-10-18 08:46 2

Introduction II: Genetic approaches to study host-pathogen interactions

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Cellular host, and restriction factors

Viruses possess small genomes that contain the bare minimum of gene­tic information to sustain replication in their host. For a successful life-cycle, viruses depend on proteins provided by the infected host cell for critical steps such as membrane attachment, genome release, replication, and egress from the host cell. Throughout the years, multiple strategies have been developed to study which host genes are important for virus infection; i.e. genes that aid the virus (host factors 2 1–3), and genes that actively hamper the virus (restriction factors 4–6). Studying these host-pathogen interactions has helped to understand how viruses acquire resistance to anti-viral treatments, what determines their tropism, how they can escape their environmental reservoirs (e.g. zoonosis), and how spreading is prevented. Furthermore, understanding the biology of virus infection also provides a dynamic and original view on cell biology of the host. Notably, much of our current knowledge of the endocytic pathway stems from studying viruses and bacteria entering the cell 7. Recent technical improvements enabling loss-of-function genetics in human cells such as RNA interference, the CRISPR-Cas9 system, and the use of haploid human cells, have greatly increased our ability to study the biology of virus infection 3,8,9. Here we will briefly explore the historical significance of host-pathogen studies as well as the recent developments in the use of genetics to study host-pathogen interactions, with an emphasis on genetic screens in haploid human cells.

Hunting host factors, one gene at a time

Host factors that are important for viral infections can be identified in various ways such as biochemical approaches (which can identify host proteins that bind to viral components), population genetics (characterizing individuals that are refractory to viral infection), or experimental genetics. Complementation or overexpression screens are a powerful example of gain- of-function genetics that yielded important insights in virology. The basis for the complementation approach is the recognition that there are cell types that are insensitive to the pathogen of interest. This often involves cells of a particular tissue or animal species. cDNA libraries isolated from susceptible cell lines can subsequently be expressed (in a clonal fashion) in non-susceptible cells enabling the selection of cDNAs that confer virus susceptibility. This method has been tremendously helpful for the identification of entry receptors such as the poliovirus receptor CD155 in

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1986 10 and the entry receptors for EV-A71; SCARB211 and PSGL-1 in 2009 12. Overexpression screens can also be applied to identify restriction factors. For instance, reporter assays in combination with the overexpression of a cDNA library in K567 cells lead to the identification of the retinoic acid inducible gene I (RIG1), a protein that is capable of activating downstream targets such as NF-kappaB and IRF-3 in presence of double stranded (ds)RNA, a well-known indicator of virus infection 13. Recently, 389 interferon-stimulated genes (ISGs) were overexpressed leading to the identification of multiple genes involved in the anti-viral innate immune response against different medically important viruses14,15 . Although the above-mentioned examples underline the strength of complemen­ tation/overexpression screens, there are limitations. The cells used for the experiments need to be non-permissible as with the receptor screens for the host factors to be detected by overexpression of their respective cDNA. The technique is also limited to the genes that are properly expressed (and therefore represented in the library), non-toxic when overexpressed, and of which the cDNA products can be cloned in the expression vectors used for the library. Furthermore, reversion of susceptibility needs to be caused by a single gene only in order to be detected with this approach 1. Finally, complementation screens usually lead to the identification of only a single host factor gene per experiment, and therefore do not generate an extensive map of the required host factors.

Going Genome-wide: Genetic tools for the identification of host factors

The technique described above can be classified as a gain-of-function genetic approach where phenotypes are associated to an increased activity of a gene of interest. This can confer virus susceptibility traits of resilient cells and thereby enables one to recover individual genes whose absence leads to resistance. The opposite loss-of-function approach would theoretically be suitable to identify all host factors that a virus requires in a susceptible cell type, yet the application of a loss-of-function approach has been limited by the availability of proper tools to inactivate genes in human cells. Nevertheless, various technological developments that we will describe below have enabled loss-of-function genetics in human cells.

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RNA interference RNA interference (RNAi) is a naturally occurring process during which small RNA duplexes (21-23 nucleotides long; guide ) lead to posttranscriptional gene silencing. These short RNA molecules are loaded on the RNA-induced silencing complex (RISC). RISC subsequently uses the guide RNA to target mRNA molecules in a sequence specific manner. Next, the targeted mRNA is digested by the endonuclease argonaute-2 (AGO2) component of RISC, leading to the reduction of the mRNA levels in the cell 8,16. The finding that these siRNAs were 2 functional in mammalian cells 17 very quickly led to the implementation of RNAi for functional genetic screens 18–21. The use of RNAi allows for an unbiased approach in a range of different cell types, and even whole organisms, and can therefore contribute to an in depth understanding of the replication cycle of many relevant viruses such as influenza viruses, rhinoviruses, and human immunodeficiency virus (HIV). The partial knockdown that is often achieved, may enable the identification of host factors that are essential for the host (in contrary to complete disruption of expression of a transcript as is the case with other approaches). Furthermore, since the inhibition occurs on mRNA level, issues with ploidy, such as with retroviral gene- trap-, chemical-, or radiation-based mutagenesis, are no longer a constraining factor. The first host-pathogen screen was performed inDrosophila with a library targeting ~91% of the predicted genes in the genome 22. The pathogen studied, Drosophila C virus (DCV), is a picornavirus known to infect insect cells. This screen identified multiple host factors (ribosomal components) required for IRES mediated translation 37 and host factors important for the formation of replication vesicles 38, highlighting the power of this new genetic approach. The applicability was further demonstrated by validating the identified genes as pan-picornavirus host factors with poliovirus infection assays in human cells 23,24. Similar studies in Drosophila were carried out using dengue virus (DENV) 25 as well as a modified influenza strain 26. Subsequent screens independently performed in human cells with HIV allowed for a reproducibility test. Four screens 27–30 were performed in different human cell lines (HeLa, 293, and Jurkat T-cells) and with different serotypes of HIV (HIV1-IIIB, HIV1-HXB2, HIV1 pseudo-typed with VSV Glycoprotein (VSV-G), and HIV1- NL4-3). The studies identified multiple host factors important for HIV infection (Brass et al.; 273 genes 27, Zhou et al.; 224 genes 28, Konig et al.; 295 genes 29, and Yeung et al.; 252 genes 30), and although the pathways were similar, the overlap of individual genes was minimal. A similar problem was observed for influenza virus screens that were carried out in the same cell type with the same strain of virus 26,31–34. A reason

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for these discrepancies could be the divergence in methodology used between the groups, but another reason might be the large amounts of false-positive and false- negative results (e.g. due to the inability to completely abolish gene expression) that often arise in RNAi based screening systems30,35,36 . Using unbiased approaches does not only lead to the identification of genes that are required for proper infection, it also unveils some aspects of the immune response of the invaded cells. RNAi proved to be very useful for the identification of important restriction factors such as IFITM3. IFITM3 was previously described to be a interferon responsive gene 37, but a siRNA screen by Brass and co-workers identified it as one of four genes (together withPUSL1 , TPST1, and WDR33) of which knock-down incited a higher influenza infection rate in U2OS cells 31. They subsequently demonstrated that the paralogs IFITM1 and IFITM2 function in a similar manner. These factors turned out to be important restriction factors for other classes of viruses as well such as DENV and West Nile virus 31. The above- mentioned examples of RNAi screens clearly demonstrate the usefulness of this approach. However, the prevalence of false positive38 and negative39 hits needs to be kept in mind.

CRISPR-Cas screens The application of targeted nucleases to screen for host factors is a relatively new approach, which became feasible with the discovery of a bacterial adaptive defense mechanism 40,41. It involves clustered regularly interspaced short palindromic repeat (CRISPR) sequences found in genomes of multiple bacterial and archaeal species 41– 43. The “random” spacers between these repeats are short fragments of foreign genomic material of previous encountered bacteriophages and plasmids 44. The spacers serve as a template for a CRISPR associated protein (Cas) to activate and guide it to the target DNA where the Cas can induce double stranded breaks with its endonuclease activity 45–47. This bacterial immune system has been adopted as a genome engineering tool applicable in eukaryotic cells and, in contrast to RNAi techniques, leads to a complete loss of expression of the targeted gene 48–50. The double stranded DNA lesions prompt non-homologous end joining (NHEJ), often resulting in deletions and frameshift mutations. This relative simple and efficient targeting of the genome allows for a rapid screening approach in a diploid cell system 51,52. Multiple libraries are available that use expression vectors for the guides and Cas proteins, which can be transduced in the desired cell line 3. Indeed, multiple host factor screens have been published (studying for example influenza virus53,54 , hepatitis C virus 55, West Nile virus 56,57), and many more viruses are expected to be studied with this exciting new genetic tool.

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Genetic screens in haploid cells Although mutagenesis-based genetic approaches in haploid organisms are not novel, this approach was never applied in human cells due to their diploid nature 58. The observation of near-haploid cells derived from a male CML patient in 1987 59 suggested that a haploid state could be compatible with the survival of somatic human cells. Cochran and coworkers subsequently subcloned these KBM-7 cells yielding a near-haploid cell line that expressed the BRC-ABL oncogene and was haploid for all chromosomes with the exception of chromosome 8. It was suggested 2 that the unique characteristics of this cell line allows for a yeast-like mutagenic- based screening approach, because disruption of a single copy of a gene would be sufficient to generate a complete knock-out of the gene-product 60. Indeed, Carette et al. transduced these cells with a retroviral gene-trap vector that would inactivate genes upon insertion and showed that a library could be created containing knockout alleles for most non-essential genes 61. This approach was applied to a phenotypic selection screen using influenza A virus (H1N1) and different bacterial toxins, identifying novel host factors for these pathogens 61. These first haploid screens were laborious: to identify the location of the gene-trap sequence, virus resistant clones had to be individually selected, cultured, isolated, and the gene-trap insertion sites had to be detected. These limitations did not allow for a large-scale sequence analysis of cells selected for phenotypes. In addition, KBM7 cells were not permissive to many virus species, making the use of these cells suboptimal for the study of a large set of viral families. Other haploid cell lines were isolated from mice (ES cells) 62–64, rats (ES cells) 65, horses (melanoma cells) 66, monkeys (ES cells) 67, and humans (ES cells) 68. Some of these cells were used for genetic approaches. Unfortunately, technical issues limited the applicability of these cell lines 63,68. After the first screens in near-haploid KBM-7 cells several technical improvements were made 69,70. An important improvement was the application of deep sequencing to examine millions of mutated genes in parallel, which enabled the detection of genes that affected virus susceptibility with high significance in a single experiment 69,70. The decision to not to select the cells for the expression of a selection marker embedded in the gene-trap vector lead to a further increase in complexity of the generated library. The power of these improvements of haploid genetics was first exemplified with four comparative host factor screens for related bacterial toxins in KBM7 cells 69. Another improvement was the generation of a different cell type. In an attempt to generate haploid induced pluripotent stem (iPS) cells, KBM7 cells were transduced with the Yamanaka transcription factors (KLF4, c-MYC, SOX2, and OCT4)71. This indeed led to the generation of iPS cells, but they did

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not remain haploid, making the cells less useful for further screening applications 71. Nonetheless, the experiment did yield a useful cell line that was only partially reprogrammed, termed HAP1. HAP1 cells are advantageous for several reasons. They are adherent, do not express any hematopoietic markers, and have lost the second copy of chromosome 8 (HAP1 cells are fully haploid with the exception of a duplicated region of chromosome 15 on chromosome 19, the derived eHAP1 cells are completely haploid 72). Importantly, HAP1 cells are, in contrast to KBM7 cells, susceptible to a variety of different viruses including vesicular stomatitis virus (VSV), and VSV virions pseudo-typed with glycoproteins from a range of different viruses. Therefore, as a first application of the use of HAP1 cells to study viral host factors, mutagenized HAP1 cells were infected with VSV pseudo-typed with Ebola virus glycoprotein. This positive selection screen led to the identification of all the members of the homotypic fusion and vacuole protein-sorting (HOPS) complex 73 (VPS11, VPS16, VPS18, VPS33A, VPS39, and VPS41) as novel entry factors for Ebola virus 73. The screen also identified Niemann-Pick 1NPC1 ( ) as a previously unknown intracellular Ebola virus receptor. The requirement of these host factors was confirmed in other cell lines with authentic Ebola and Marburg virus. Furthermore, mice deficient forNPC1 were shown to be resistant to Ebola and Marburg virus infection 73, underscoring the validity of this unbiased, genome wide approach and HAP1 cells as a model system for studying the entry of viruses. To date, HAP1 cells have proven to be permissive for a wide range of unrelated viruses including different picornaviruses74–76 , DENV 55, adeno associated virus (AAV) 77, and many other virus species of which an overview is provided in Table 1, enabling an extensive exploration of host-pathogen interactions in human cells. A limitation of the classic loss-of-function screens performed in HAP1 cells is the stringent selection method. Most studies are performed using cytopathic viruses, under conditions where that vast majority of the mutagenized cell population die in order to select for mutants that survive. Therefore, this approach is biased towards host factors that create a strong resistance phenotype upon mutation and it can only be used for viruses that are cytotoxic. Pillay et al. presented a creative solution to this problem to study host factor requirements of the non-cytopathic AAV2. They infected a HAP1 mutagenized library with an AAV2 which is genetically engineered to express red fluorescent protein (RFP), then sorted the cells (by using fluorescence-activated cell sorting (FACS)) from the infected cell population that were low in RFP expression. By doing so, they were able to identify host-factors, including a novel entry receptor (KIAA0319L, or AAVR), critical for the infection of most AAV serotypes. Furthermore, they were able to provide evidence that

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Virus Cell Type Host-factors Reference

Influenza virus (H1N1) KBM7 Sialic acid biosynthesis Carette et al., 2009 61 (CMAS, SLC35A2) Ebolavirus (VSV pseudo-typed) HAP1 HOPS-complex, NPC1 Carette et al., 2011 73 Lassa virus (VSV pseudo-typed) HAP1 DAG1, LAMP1 Jae et al., 2013 78 Kaposi's sarcoma-associated KBM7 PLP2 Timms et al., 2013 79 herpesvirus Andes virus (VSV pseudo-typed) HAP1 MBTPS1, MBTPS2, SCAP, and Petersen et al., 2014 80 2 SREBF2 Hanta virus (VSV pseudo-typed) HAP1 SREBF2, MBTPS1, MBTPS2, Kleinfelter et al., 2015 81 SCAP Rift Valley fever virus HAP1 Heparan sulfate, COG-complex Riblett et al., 2015 82 Enterovirus D68 HAP1 Sialic acid biosynthesis, Baggen et al., 2016 76 PLA2G16 Dengue virus HAP1 OST complex, TRAP complex, Marceau et al., 2016 55 EMC, ERAD Adeno-associated virus HAP1 AAVR, GARP-complex Pillay et al., 2016 77 Poliovirus type 1 HAP1 PVR, PLA2G16 Staring et al., 2017 74 Coxsackievirus B1 HAP1 CAR, PLA2G16 Staring et al., 2017 74 Coxsackievirus B3 HAP1 CAR, PLA2G16 Staring et al., 2017 74 Coxsackievirus A7 HAP1 LDLR, SCARB2, PLA2G16 Staring et al., 2017 74 Enterovirus A71 HAP1 SCARB2, PLA2G16 Staring et al., 2017 74 Lujo Virus (VSV pseudo-typed) HAP1 NRP2, CD63 Raaben et al., 2017 83 virus HAP1 GARP-complex, COG-complex, Realegeno et al., 2017 84 Heparan sulfate Thrombocytopenia syndrome HAP1 UGCG Drake et al., 2017 85 virus (VSV pseudo-typed) Coxsackievirus A10 HAP1 KREMEN1, PLA2G16, PI4K, Staring et al., 2018 75 C10ORF76 Human endogenous HAP1 Heparan sulfate Robinson-McCarthy et al., (VSV pseudo-typed) 2018 86

Table 1. Forward genetic screens performed in human haploid cells to identify host/virus interactions.

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AAVR is important for virus entry in a range of different cell lines as well as in anin vivo model for AAV gene-therapy 77. As described in chapter 3, a similar FACS based approach can also be applied to study restriction factors. To conclude, the use of genetic perturbation screens has proven to be a very successful method to gain more insight in host-pathogen interactions. The next three experimental chapters in this thesis provide examples of the application of haploid genetics to study the entry of the picornaviruses.

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80. Petersen, J. et al. The Major Cellular Sterol Regulatory Pathway Is Required for Andes Virus Infection. PLoS Pathog. 10, (2014). 81. Kleinfelter, L. M. et al. Haploid genetic screen reveals a profound and direct dependence on cholesterol for hantavirus membrane fusion. MBio 6, (2015). 82. Riblett, A. M.et al. A Haploid Genetic Screen Identifies Heparan Sulfate Proteoglycans Supporting Rift Valley Fever Virus Infection.J. Virol. 90, 1414–23 (2015). 83. Raaben, M. et al. NRP2 and CD63 Are Host Factors for Lujo Virus Cell Entry. Cell Host Microbe 22, 688–696 (2017). 84. Realegeno, S. et al. Host Factor Screen Using Haploid Cells Identifies Essential 2 Role of GARP Complex in Extracellular Virus Formation. J. Virol. 91, e00011-17 (2017). 85. Drake, M. J. et al. A role for glycolipid biosynthesis in severe fever with thrombocytopenia syndrome virus entry. PLoS Pathog. 13, (2017). 86. Robinson-McCarthy, L. R. et al. Reconstruction of the cell entry pathway of an extinct virus. bioRxiv (2018). 87. Hoffmann, H.-H.et al. Diverse Viruses Require the Calcium Transporter SPCA1 for Maturation and Spread. Cell Host Microbe 22, 460–470.e5 (2017).

PS_JS_def.indd 41 08-10-18 08:46 go all the way. it could mean not eating for 3 or 4 days. it could mean freezing on a park bench. it could mean jail, it could mean derision, mockery, isolation.

PS_JS_def.indd 42 08-10-18 08:46 3

PLA2G16 represents a switch between entry and clearance of Picornaviridae

Authors: Jacqueline Staring1, Eleonore von Castelmur1, Vincent A. Blomen1, Lisa G. van den Hengel1, Markus Brockmann1, Jim Baggen2, Hendrik Jan Thibaut2, Joppe Nieuwenhuis1, Hans Janssen1, Frank J.M. van Kuppeveld2, Anastassis Perrakis1, Jan E. Carette3 and Thijn R. Brummelkamp1, 4, 5

Affiliations: 1Netherlands Cancer Institute, Plesmanlaan 121, 1066 CX, Amsterdam, The Netherlands.

2Virology Division, Department of Infectious Diseases and Immunology, Faculty of Veterinary Medicine, Utrecht University, Utrecht, The Netherlands.

3Department of Microbiology and Immunology, Stanford University School of Medicine, 299 Campus Drive, Stanford, CA 94305.

4CeMM Research Center for Molecular Medicine of the Austrian Academy of Sciences, 1090 Vienna, Austria.

5Cancer GenomiCs.nl (CGC.nl), Plesmanlaan 121, 1066 CX Amsterdam, The Netherlands.

Nature 2017 541, 412-416

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Picornaviruses are a leading cause of human and veterinary infections that result in various diseases, including polio and the common cold. As archetypical non-enveloped viruses, their biology has been extensively studied1. Although a range of different cellsurface receptors are bound by different picornaviruses2–7, it is unclear whether common host factors are needed for them to reach the cytoplasm. Using genome-wide haploid genetic screens, here we identify the lipid-modifying enzyme PLA2G16 (refs 8–11) as a picornavirus host factor that is required for a previously unknown event in the viral life cycle. We find that PLA2G16 functions early during infection, enabling virion-mediated genome delivery into the cytoplasm, but not in any virion-assigned step, such as cell binding, endosomal trafficking or pore formation. To resolve this paradox, we screened for suppressors of the ∆PLA2G16 phenotype and identified a mechanism previously implicated in the clearance of intracellular bacteria12. The sensor of this mechanism, galectin-8 (encoded by LGALS8), detects permeated 3 endosomes and marks them for autophagic degradation, whereas PLA2G16 facilitates viral genome translocation and prevents clearance. This study uncovers two competing processes triggered by virus entry: activation of a pore-activated clearance pathway and recruitment of a phospholipase to enable genome release.

To identify picornavirus host factors, mutagenized HAP1 cells were infected with different picornaviruses. Gene-trap insertions mapped in resistant cell populations (Fig. 1a) were enriched for mutations in the respective entry receptors such as the poliovirus receptor (PVR, n = 475 independent mutations) (Fig. 1b–e, Extended Data Fig. 1a and Supplementary Tables 1–5). In addition, PLA2G16, a gene not linked to virus infection before, was identified as a common host factor.PLA2G16 encodes a small phospholipase implicated in obesity13 located on the same genomic locus as its homologues RARRES3 and HRASLS2, which were not enriched for disruptive mutations (Extended Data Fig. 1b). In line with the absence of PLA2G16 in previous haploid screens (performed with other viruses)14–18, cells carrying a gene-trap insertion in the PLA2G16 locus specifically displayed resistance to picornaviruses (Fig. 1f). Resistance to virus infection could be overcome by expression of a PLA2G16 cDNA, which required the catalytic site cysteine (Cys113) (Extended Data Fig. 1c), suggesting that enzymatically active PLA2G16 functions as an important host factor for infection. PLA2G16 inactivation using CRISPR/Cas9 gene-editing in HAP1, HeLa, HEK293T and A549 cells led to poliovirus type 1 (PV1) resistance (Extended Data Fig. 1d, e and Supplementary Table 7), and deficiency ofPLA2G16 in HeLa

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F igure 1. Identification of PLA2G16 as a host factor for picornaviruses. a, Schematic overview of mutagenesis-based screen in haploid cells. b–e, Positive selection screens for picornavirus infection: PV1 (strain Sabin) (b) coxsackievirus B1 (CV-B1) (c), CV-A7 (d) and enterovirus A71 (EV-A71) (e). Each circle represents a gene, with size corresponding to the number of insertion sites per gene. They axis indicates the significance of enrichment of several insertions in the affected gene compared to an unselected control. Genes were randomly distributed on the x axis. Coloured circles indicate known virus receptors and PLA2G16. f, Infection of wild-type (HAP1) and PLA2G16 gene-trap insertion (PLA2G16-GT) cells with different viruses. Viable cells were stained using crystal violet (n = 3). g, Wild-type, ∆PLA2G16 and ∆PLA2G16 cells expressing

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PLA2G16 cDNA were infected with picornaviruses. Cells were stained 8 h after infection with a double-stranded RNA (dsRNA) antibody, and infection efficiencies were quantified (mean± s.d., n = 4, chi-square test, ****P < 0.0001). h, dsRNA staining of muscle tissue (tongue) of CV-A10 virus infection in wild-type and ∆Pla2g16 mice. i, Kaplan–Meier graphs of infected wild-type and ∆Pla2g16 mice exposed to CV-A10 (left) and CV-A4 (right).P < 0.0001 (log-rank test) for survival of ∆Pla2g16 mice compared to wild type for both CV-A10 and CV-A4.

cells resulted in resistance to enteroviruses (including rhinoviruses) as well as picornaviruses of the Cardiovirus genus (encephalomyocarditis virus (EMCV), Saffold virus) (Fig. 1g and Extended Data Fig. 1f). We then generated Pla2g16-deficient mice (Extended Data Fig. 2a, b), previously reported to be resistant to diet-induced obesity13 (Extended Data Fig. 2c, d). Fibroblasts from these ∆Pla2g16 mice showed resistance to infection by coxsackievirus A10 (CV-A10, Extended Data Fig. 2e). Injecting 1-day-old pups 3 intraperitoneally with CV-A10 (n = 14 wild type, n = 12 ∆Pla2g16) or CV-A4 (n = 12 wild type, n = 8 ∆Pla2g16) resulted in high levels of double-stranded viral RNA staining in muscle tissue; whereas only modest staining was observed in Pla2g16- knockout mice (Fig. 1h). Notably, although wild-type mice succumbed to infection and displayed symptoms such as paralysis (Fig. 1i), most ∆Pla2g16 animals infected with CV-A10 or CV-A4 (9 out of 12, or 12 out of 12 pups, respectively) showed no signs of illness for the duration of the experiment (at least 18 days). To study the role of PLA2G16 in the infection cycle, we used fluorescently labelled poliovirus (Cy5-PV1). ∆PLA2G16 HeLa cells showed normal levels of plasma membrane binding and subsequent internalization, indicating that PLA2G16 is not required for receptor binding or uptake into cells (Extended Data Fig. 3a, b). Infection of ∆PLA2G16 cells complemented with a green fluore- scent protein (GFP)-tagged PLA2G16 transgene resulted in the appearance of cytoplasmic foci. These foci were in proximity to the Cy5-PV1 signal (Fig. 2a and Supplementary Video 1) and their appearance was prevented by the capsid-binding (Extended Data Fig. 3c). Given the proximity of the lipid- modifying enzyme PLA2G16 to incoming virions, we asked whether PLA2G16 functions in pore formation—the next step of the infection cycle. Virus-induced pore formation enables the cellular uptake of the proteinaceous 19 ribosome-inactivating toxin α-sarcin (Fig. 2b). Notably, exposure of either wild- type or ∆PLA2G16 HeLa cells to PV1 in combination with α-sarcin readily inhibits translation in both cell lines (Fig. 2c), indicating that PLA2G16 is not essential for membrane permeabilization.

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F igure 2. PLA2G16 functions after entry and before replication. a, Localization of GFP–PLA2G16 (green) and Cy5-labelled PV1 (red; multiplicity of infection (MOI) value of ∼10) in live HeLa cells 15 min after infection. Individual channels for different regions are shown. b, c, Schematic overview of pore-formation assay using α-sarcin (b), and a pore-formation assay with co-incubation of PV1 and α-sarcin (c). Translation was measured using 35S-methionine/ cysteine-labelled incorporation. α-Tubulin served as loading control. For gel source data, see Supplementary Fig. 1. WT, wild type. d, Infection or transfection of PV1-DsRed (red) in the indicated genotypes. Cells were transfected with the viral genome (RNA) and DsRed expression was compared to expression in cells infected with PV1-DsRed virions. Cells were counterstained with Hoechst33342 (blue) for DNA. Inhibition of replication using GHL (2 mM) indicated that viral replication was required for the detection of DsRed expression (percentage values denote mean ± s.d., n = 3).

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Picornavirus replication involves the accumulation of intracellular membrane structures, which are used for RNA replication. ∆PLA2G16 cells rarely contained these viral replication sites (Extended Data Fig. 3d), suggesting that PLA2G16 acts after pore formation but before or concurrently with replication. To address this, we infected wildtype cells with a DsRed-expressing PV1. This virus must replicate to produce detectable fluorescence. Although many wild-type cells showed fluorescence (Fig. 2d) that could be blocked with the replication inhibitor guanidine hydrochloride (GHL)20, no fluorescence was observed after infection of ∆PLA2G16 cells. However, direct transfection of the viral genome into the cells bypassed the requirement for PLA2G16, and led to the generation of viral progeny as in wild-type cells (Fig. 2d and Extended Data Fig. 3e, f), indicating that PLA2G16 is not required for viral RNA replication. Collectively, our data point to a role for PLA2G16 in delivery of the viral genome to the cytoplasm from virions. 3 To gain additional insight into the phenotype of ∆PLA2G16 cells, we designed a genetic suppressor screen to find mutants restoring virus susceptibility in ∆PLA2G16 cells. We exposed mutagenized ∆PLA2G16 HAP1 cells to PV1-DsRed, and used flow cytometry to select populations of DsRed-positive and -negative cells (Fig. 3a). For genes not affecting susceptibility to virus infection, gene-trap mutations should be recovered with comparable frequencies in both populations. However, the recovery of mutations in factors that affect virus susceptibility should be biased. Indeed, mutations in PVR were enriched in the DsRed-negative cell population (Extended Data Fig. 4a). Notably, in the DsRed-positive cell population, we identified a large number of genes involved in autophagy, membrane trafficking and glycosylation (Fig. 3b and Supplementary Table 6). This included the danger receptor LGALS8 (also known as galectin-8), which was previously linked to autophagic clearance of intracellular bacteria12. This cytoplasmic lectin binds glycans of the external leaflet of host membranes, which become exposed when bacteria generate pores in the vacuole in which they reside, leading to the formation of autophagosomes. To determine whether a similar mechanism may affect viral infection, we expressed a GFP-tagged galectin-8 transgene (GFP–LGALS8) in ∆PLA2G16 cells and exposed them to wild-type PV1 or coxsackievirus B1 (CV- B1). This led to the formation of cytoplasmic GFP–LGALS8 foci within minutes after infection (Extended Data Fig. 4b). Although the related galectin-3 (encoded by LGALS3) is recruited by adenovirus-induced membrane rupture, this has not been linked to viral restriction21. To test whether LGALS8 affects viral infection, we generated HeLa cells carrying single or double mutations in PLA2G16, LGALS8 or the essential autophagy factor ATG7 (Extended Data Fig. 4c). Exposure of these

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F igure 3. A suppressor screen identifies a pore-based restriction mechanism in∆ PLA2G16 cells. a, Schematic overview of PV1-DsRed suppressor screen using flow cytometric cell sorting of mutagenized haploid ∆PLA2G16 cells. b, PV1-DsRed suppressor screen in ∆PLA2G16 cells. Each

dot represents a gene positioned on the x axis by the total amount of insertions per gene (log10). The y axis represents the mutation index (the relative number of integrations in the DsRed-high population divided by the relative number of integrations in the DsRed-low population for each individual gene). c, d, Galectin-8 (LGALS8) deficiency c( ) or ATG7 deficiency d( ) restores virus susceptibility in ∆PLA2G16 HeLa cells. Indicated genotypes were infected with an increasing amount of PV1 or CV-B3–GFP, and infected cells were detected using a dsRNA antibody or by measuring GFP expression, respectively (n = 4). Cells were counterstained with Hoechst33342 (blue) for DNA.

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3

F igure 4. PLA2G16 facilitates genome dislocation from LGALS8 clusters and enables viral genome translation. a, Wild-type and ∆PLA2G16 HeLa cells expressing GFP-tagged galectin-8 (GFP–LGALS8; green) infected with PV1 and CV-B1 (MOI of ∼20). Hypotonic shock serves to induce membrane damage. Insets represent magnifications of dashed boxes.b , Schematic overview of LGALS8 autophagic flux assay. c, Western blot analysis of autophagy-dependent GFP–LGALS8 cleavage during hypotonic shock (top) and infection with PV1 (middle) or CV-B1 (bottom) in the indicated genotypes. m.p.i., minutes post infection. d, In situ hybridization to detect PV1 RNA genomes (red) 30 min after infection in wild-type and ∆PLA2G16 cells expressing GFP–LGALS8 (green). Circles represent examples of LGALS8 foci. In the pie charts (bottom), grey denotes LGALS8 foci positive for RNA, black denotes LGALS8 foci negative for RNA. ****P < 0.0001, chi-square test, n = 3. e, Translation of incoming viral RNA using a CV-B3 expressing Renilla luciferase (CV-B3-luc). GHL served to block viral genome replication. **P < 0.01, ***P < 0.001, unpaired two-sided Welch-corrected t-test, n = 3. RLU, relative light units. f, Transfection of CV-B3-luc RNA in wild-type versus ∆PLA2G16 cells. n = 3; NS, not significant.g , Protein lysates of CV-B1-infected cells (MOI = 10, 2 h after infection) probed for eIF4G in indicated genotypes. GHL was added 30 min before infection. For gel source data, see Supplementary Fig. 1. h, Schematic model of the role of PLA2G16 during picornavirus entry.

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cells to PV1, CV-B1 or CV-B3–GFP demonstrated that loss of either LGALS8 or ATG7 restored virus susceptibility in ∆PLA2G16 cells (Fig. 3c, d and Extended Data Figs 4d–f, 5a–c), suggesting that their resistance phenotype is at least in part a result of virus-induced LGALS8 recruitment. As LGALS8 and PLA2G16 showed a genetic interaction, we investigated their connection. PLA2G16 deficiency did not affectLGALS8 recruitment by infection or pathogen-free (sterile) membrane damage induced by hypotonic shock12 (Fig. 4a). Mutation of the carbohydrate-recognition domains (CRDs) of LGALS8 suggested that binding of host glycans to the N-terminal CRD trigger virus-induced LGALS8 recruitment, similar to bacterial pathogens12 (Extended Data Fig. 6a, b). As PLA2G16 formed foci in infected cells (Fig. 2a), we asked whether PLA2G16 responded to membrane damage. Notably, as observed for LGALS8, GFP– PLA2G16 formed distinct cytoplasmic foci when cells were exposed to hypotonic shock (Extended Data Fig. 7a, b), and this was independent of LGALS8 (Extended Data Fig. 7c). Similarly, lysosome rupture led to the recruitment of LGALS8 (ref. 12) and subsequent accumulation of GFP–PLA2G16 (Extended Data Fig. 8a–d). Importantly, LGALS8 recruitment to damaged endosomes or ruptured lysosomes did not require PLA2G16 and vice versa (Extended Data Fig. 8e, f). Also, the localization of PLA2G16 to incoming viruses was independent of LGALS8 (Extended Data Fig. 8g). We found that the hydrophobic C terminus of PLA2G16 is needed to respond to membrane damage, suggesting that PLA2G16 may detect membrane damage autonomously (Extended Data Fig. 8h). To determine whether PLA2G16 affects LGALS8 function, we modified an assay to measure autophagic flux involving release of GFP from the autophagy protein LC3-GFP by lysosomal proteases22,23 (Fig. 4b). Exposure of GFP–LGALS8- expressing cells to hypotonic shock, PV1 or CV-B1 resulted in the formation of LGALS8 foci (Fig. 4a and Extended Data Fig. 9a) and increased the amount of free GFP (Fig. 4c), which required the autophagic machinery. GFP cleavage occurred to the same extent in ∆PLA2G16 cells (Fig. 4c), suggesting that the ability of LGALS8 to trigger proteolysis of associated molecules is not influenced by PLA2G16. Because LGALS8 activity was unaffected by PLA2G16, we followed the fate of the viral genome using LGALS8 as a tool to mark virions that have formed pores. In wild-type and ∆PLA2G16 cells, viral genomes were initially detected at the plasma membrane (Extended Data Fig. 9b), which internalized at later time points (Extended Data Fig. 9c). In wild-type cells, most GFP–LGALS8 foci did not contain viral genomes (Fig. 4d). The opposite was observed in ∆PLA2G16 cells, in which most LGALS8 foci did contain viral genomes. Crucially, this suggests

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that PLA2G16 facilitates the displacement of the viral genome from LGALS8- positive vesicles, which may promote viral translation. To test this, we measured viral translation before replication. Indeed, ∆PLA2G16 cells displayed decreased translation of a viral luciferase transgene upon infection, regardless of replication inhibition (Fig. 4e), but, consistent with previous data, not upon transfection (Fig. 4f). As an indicator of viral protein production, we also measured the cleavage of eIF4G by 2A protease in infected cells. Consistently, eIF4G cleavage was attenuated in ∆PLA2G16 cells even when virus replication was blocked (Fig. 4g and Extended Data Fig. 9d). Together, this suggests that PLA2G16 does not affect the detection or autophagic clearance of permeated endosomes by LGALS8, but stimulates cytoplasmic availability of the viral genome to enable viral protein synthesis and subsequent replication (Fig. 4h). In conclusion, we describe two membrane-linked mechanisms with opposing roles during picornavirus infection. The relocalization of PLA2G16 after membrane 3 perturbation suggests that it may act as a cellular sensor of membrane damage at sites of virus entry. Its function may resemble a strategy used by parvoviruses 24,25 carrying a capsid-linked PLA2 to enable endosomal escape . Similarly, PLA2G16 could be involved in initiating pore formation, increasing pore size, or in maintaining pores for genome delivery. Alternatively, modifying the membrane to which the capsid is anchored may alter membrane curvature and create forces towards the RNA-containing capsid, facilitating genome release. Remarkably, the 2A proteins of certain picornaviruses (for example, Aichivirus, parechovirus, and avian encephalomyelitis) are homologues of PLA2G16 (ref. 26). This similarity, noted years ago, may now be related to a role for cellular PLA2G16 function in viral infection. We also demonstrate how membrane damage can trigger viral restriction. Viral restriction poses an intriguing impossibility: effective restriction mechanisms cannot exist; otherwise the affected virus would not exist. Thus, typically, restriction mechanisms are inactive in the host or cell type in which the virus replicates, or counteracted by viral factors. Ironically, however, here a cellular factor counters the restriction mechanism, posing intriguing questions on the co-evolution of these two processes.

Online Content Methods, along with any additional Extended Data display items and Source Data, are available in the online version of the paper; references unique to these sections appear only in the online paper.

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Methods

No statistical methods were used to predetermine sample size. The experiments were not randomized.

Cells and culture conditions HAP1 cells were generated from KBM7 cells as described before15. H1-HeLa, HEK293T and A549 cells were obtained from ATCC. Mouse-tail fibroblasts were obtained from C57BL/6N wild-type and ∆Pla2g16 mice. HAP1 cells were cultured in IMDM (ThermoFisher Scientific) supplemented with 10% fetal calf serum (FCS), penicillin–streptomycin and l-glutamine (ThermoFisher Scientific). H1- HeLa, HEK293T, A549 cells, and mouse-tail fibroblasts were cultured in DMEM (ThermoFisher Scientific) supplemented with 10% FCS, penicillin–streptomycin and l-glutamine. Cell lines were obtained from authentic stocks (ATCC), or authenticated by genotyping. All cell lines were tested for mycoplasma and were verified to be negative.

Viruses The following picornaviruses were used: PV1 (strain Sabin, ATCC), poliovirus- DsRed type 1 (strain Mahoney, obtained by transfection of the infectious clone pPVM-2A50-DsRed as described before27), coxsackievirus B1 (strain Conn-5, ATCC), CV-B3 (strain Nancy, obtained by transfection of the infectious clone p53CB3/T7 as described before28, CV-B3-Renilla luciferase (strain Nancy, was obtained by transfection of the infectious clone pRLuc-53CB3/T7 as described previously28), CV-B3–GFP (strain Nancy, was obtained by transfection as described previously29), CV-A4 (strain H08302 509, national collection of pathogenic viruses (NCPV)), CV-A5 (strain Swartz, ATCC), CV-A7 (strain Parker, ATCC), CV-A10 (strain Kowalik, ATCC), rhinovirus 2 (strain HGP, ATCC), enterovirus A71 (strain BrCr, ATCC), echovirus 7 (strain Wallace, ATCC), and EMCV (VR-1762, ATCC), Saffold virus 3 (isolated from a stool sample30). All picornaviruses were propagated in H1-HeLa cells and the titre was determined by standard plaque assays. Other viruses used to determine specificity for PLA2G16 requirement were adenovirus (ATCC), virus 1 (strain KOS, provided by H. Ploegh), influenza virus (strain A/PR/8/34, Charles River), vesicular stomatitis virus (VSV), and VSV pseudotyped with glycoproteins of Ebola virus, rabies virus, and (provided by S. Whelan).

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Reagents and antibodies l-leucyl-l-leucine methyl ester (Leu-Leu-Ome, LLOme), GHL, pleconaril and protamine sulfate were purchased from Sigma- Aldrich. α-Sarcin toxin was from Santa Cruz. Restriction enzymes were purchased from New England Biolabs. Antibodies directed against α-tubulin (10D8), GFP (B-2), CDK4 (C-22) and LAMP1 (H4A3) were from Santa Cruz. Wheat germ agglutinin (WGA), Hoechst33342, Alexa Fluor 488 and Alexa Fluor 568 were purchased from ThermoFisher Scientific. Rictor (D16H6) and eIF4G antibodies were from Cell Signaling Technology. Antibody directed against PLA2G16 was purchased from Abnova, ATG7 antibody was from Abgent, J2 anti-dsRNA antibody was from English and Scientific Consulting, VP1 antibody (5-D8/1) was from Leica Biosystems and the secondary antibodies conjugated to horseradish peroxidase (HRP) were from Biorad.

Genetic screen for picornavirus host factors 3 Positive selection screens for genes required for picornaviruses were performed using HAP1 cells that have been mutagenized using a gene trap retrovirus as described previously16,31. In brief, HEK293T cells were seeded in 12 T175 flasks and transfected with packaging and gene-trap plasmids14. Virus was collected first after 48 h, twice a day for at least 3 days, and centrifuged for 2 h at 60,000g at 4 °C to concentrate. Forty million HAP1 cells were seeded in T175 flasks and infected with the gene-trap retrovirus in the presence of 8 μg ml−1 protamine sulfate. After passaging for approximately 5 days, the mutagenized cells were frozen down as libraries for genetic screening. For selections using the various picornaviruses, 100 million mutagenized HAP1 cells were seeded in 14 T175 flasks in the presence of PV1, CV-B1, CV-B3, CV-A7 and enterovirus A71, each with an MOI of approximately 5. Resistant colonies were recovered after 10 days. Cells were subsequently pooled and used for genomic DNA isolation and for sequencing of gene-trap insertion sites.

Genetic screen for suppressor mutations in ∆PLA2G16 HAP1 cells To identify suppressor mutations in a PLA2G16-deficient background, a library of gene-trap mutagenized AAV-targeted ∆PLA2G16 HAP1 cells was taken in culture and seeded in 30 T175 flasks. Upon reaching confluence, 3 × 109 cells were infected with PV1-DsRed (MOI of 5) and incubated at 37 °C for 7 h. Subsequently, cells were washed with PBS, trypsinized, pelleted, and fixed with 4% paraformaldehyde (Sigma Aldrich) in PBS for 10 min, pelleted, and subjected to an additional fixation using BD Fix Buffer I (BD Biosciences) according to manufacturer’s specification (10 min at 37 °C). After washing twice using PBS containing 10% FCS the cells

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were FACS sorted on a Biorad S3 sorter. The 1% of cells with the lowest and highest fluorescence signal of PV1-DsRed were sorted until more than 6 million cells were acquired for both gates. Per population, sorted cells were pelleted. Genomic DNA was de-crosslinked and isolated as described previously31.

Amplifying and mapping insertion sites For both types of screens the gene-trap insertion sites were amplified using a LAM- PCR procedure previously described in detail31. In short, a single biotinylated primer was used to linearly amplify the gene trap cassette and flanking genomic region for 120 cycles. For the positive selections 8 LAM-PCR reactions were used and for the suppressor screen 6 reactions were amplified in both channels. Subsequently, the resulting single stranded DNA products were captured on magnetic streptavidin- coated beads (M-270, ThermoFisher Scientific), washed, and a single-stranded DNA (ssDNA) linker was ligated to facilitate a final round of exponential amplification for 18 cycles after washing. For the linker ligation, either a 5′-phosphorylated linker was used in combination with Circligase II (Illumina) or using a purified RNA ligase in combination with a 5′-adenylated linker, described in detail here31. Subsequently, the products of final PCR amplification were purified (Qiagen, PCR purification kit) and sequenced on a HiSeq2000 (Illumina) or HiSeq2500 (Illumina) (18 pmol loading concentration) using a custom sequencing primer. For mapping the insertion sites, reads were aligned to the human genome (hg19) using bowtie32 and only reads aligning with zero or a single mismatch were selected as flanking regions to identify insertion sites. The reads were subsequently assigned to the non-overlapping regions of protein-coding genes (Refseq hg19) using intersectBed and unique insertions counted per gene. Insertions less than 2 base pairs apart were removed. For the positive-selection screens for picornavirus host factors, a similar method was used as previously reported15,16,18,33. The number of disruptive integrations (in the same orientation as the gene) was compared using a one-sided Fisher’s exact test to an unselected HAP1 control dataset previously published16 and available at the SRA (SRP018361: accession SRX223544).P values were false discovery rate (FDR)-corrected (Benjamani—Hochberg). The results of the experiments are displayed in bubble plots with the −log(significance) on they axis and the genes randomly distributed over the x axis in order to minimize overlap. For finding suppressor mutants in the∆ PLA2G16 cells, insertion sites were mapped for both the pool with the lowest as well as the highest PV1-DsRed signal independently in similar fashion, although insertions less than 2 bp apart were not removed. However, to find genes that deviate most between both populations, instead

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of comparing both to an unselected control dataset, the datasets were mutually compared using a two-sided Fisher’s exact test (P values were subsequently FDR- corrected). This was visualized by plotting the total number of unique insertions in each gene on the x axis and the mutation index (MI: the relative number of integrations in the high population divided by the relative number of integrations in the low population) on the y axis. Genes with integrations mapped in only a single channel were corrected by addition of 1 in the other channel to calculate the mutation index.

Generation of isogenic knockout cell lines Knockout cell lines were generated by using different gene-targeting approaches: transcription activator-like effector nucleases (TALENs), clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9, and integration of a gene- trap vector or adeno-associated virus (AAV) vector. All knockout cell lines are single- 3 cell clones, and gene editing was verified by Sanger sequencing (Supplementary Table 7). TALENs were designed in order to edit exon 2 in human PLA2G16 gene in HeLa cells. Subsequently, TALENs were assembled according to the TALE construction protocol adapted from ref. 34 (Addgene, kit 1000000019). In short, TALE binding domains targeting PLA2G16 consisted of two 20-nucleotide-long stretches of dsDNA with a 19-nucleotide-long spacer in between, with both stretches starting with a thymidine. Each selected DNA stretch was divided into 3 hexamers and subsequently assembled using Golden Gate digestion-ligation. Afterwards, the successfully ligated hexamers were amplified in a PCR reaction and subsequently cloned into a TALE backbone vector according to the Golden Gate digestion ligation protocol. Successful custom-made TALENs were selected for transfection in HeLa cells using Turbofectin (Origene) following the manufacturer’s protocol. Two days after transfection, HeLa cells were maintained in DMEM containing 90 μg ml−1 of blasticidin (Invivogen) and subsequently, selected for TALEN integration into target gene. Most HeLa knockout cell lines were generated using CRISPR/Cas9 gene editing technique. CRISPR guide RNAs (gRNAs) were constructed targeting PLA2G16 (exon 3; 5′-GGCCGGGAGTGACAAGTACC-3′), PVR (exon 2: 5′- TGGGCGCGGCATGGTGAATC-3′ and 5′-GATGTTCGGGTTGCGCGTAG-3′), LGALS8 (exon 2: 5′-AGTGACGCAGACAGGTAAAA-3′; exon 3: 5′-CGAGCCG ATGTGGCCTTTCA-3′) and ATG7 (exon 2: 5′-AGAAGAAGCTGAAC GAGTAT-3′), and ligated into the backbone vector pX330 (ref. 35). HeLa cells were

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transfected with CRISPR/Cas9 and a blasticidin-resistance cassette as a transfec- tion control, which was performed using Lipofectamin2000 (Invitrogen) following the manufacturer’s protocol. Blasticidin selection of HeLa cells was started 2 days after transfection. LentiCRISPR targetingPLA2G16 was used for generation of PLA2G16 knockout cell lines for Fig. 1h. HEK293T cells were transfected with lentiCRISPR (pXPR_001 vector; Zhang laboratory) expressing a PLA2G16- targeting gRNA in combination with a mix of ∆VPR, VSVg and Adv packaging plasmids, thereby using Turbofectin according to the manufacturer’s manual. Cells were subsequently transduced with PLA2G16 CRISPR lentivirus in the presence of 8 μg ml−1 protamine sulfate (Sigma). As control for PLA2G16 lentiCRISPR transduction, HAP1, HEK293T, A549 and HeLa cells were infected with a control lentiCRISPR lacking a gene-specific gRNA. Gene-trap HAP1 knockout clones were obtained by sub cloning the mutagenized PV1-selected cell population. To produce infectious recombinant human AAV virions, 293T cells were cotransfected with pAAV-MCS, pAAV-RC and pHelper (Cell Biolabs, Inc.). Recombinant AAV vector was subsequently harvested and used for transduction of HAP1 cells to target exon 3 of PLA2G16.

Constructs and plasmids PLA2G16 and N-terminally tagged PLA2G16 were amplified by PCR from cDNA (generated using SuperScript First Strand Synthesis System (Invitrogen) de- rived from HAP1 cells) using the following primers: 5′-GAATTCACCATGC- GTGCGCCCATTCCAGAG-3′ and 5′-GTCGACTTATT GCTTTTGTCGCTT- GTTTC-3′. The PCR product was digested with EcoRI and SalI, and cloned into retroviral vectors pMX-2A-Blast (digested with EcoRI and XhoI), pMX-FLAG- 2A-Blast (digested with EcoRI and XhoI), and pBabe-Puro (digested with EcoRI and SalI). N-terminally Flag-tagged mouse Pla2g16 was obtained by gene synthesis (Life Technologies) based on Ensembl transcript ENSMUST00000025925, ampli- fied by PCR (5′-GAATTCACCAT GCTAGCACCCATACCAG-3′, and 5′-GTC- GACTTATTGCTTCTGTTTCT TGTTTC-3′), digested with EcoRI and SalI, and cloned into the retroviral vector pMX-FLAG-2A-Blast (digested with EcoRI and XhoI). The humanPLA2G16 - C113A mutant construct was obtained by site-direct- ed mutagenesis using pMX- PLA2G16 as template. Human GFP-tagged PLA2G16 was obtained by gene synthesis (Life Technologies) based on Ensembl transcript ENST00000323646, amplified by PCR (5′-AATTCCAGATCTGCCACCATG-3′ and 5′-AGATCTGGACATTTTATTGCTTTTGTCGC-3′), digested with BglII, and cloned into pBabe-Puro with BamHI and SalI. Truncated GFP–PLA2G16 was

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amplified from full-lengthPLA2G16 (5′-AATTCCAGATCTGCCACCATG-3′ and 5′-AGATCTGGAATTTTAATCTCTGACCTGGTCACTGC-3′) digest- ed with BglII, and cloned into pBabe-Puro with BamHI and SalI. GFP–LGALS8 (5′-AATTCCAG ATCTATGGTGAGCAAG-3′ and 5′-GGAATTAGATCTCTAC- CAGCTCC-3′); mCherry–LGALS8 (5′-AATTCCAGATCTGCCACCATGGT- GTCC-3′ and 5′-GGAATTAGATCTCTACCAGCTCC-3′); GFP–LGALS8-R69H (5′-CATGAGATCTACCGCCACCA-3′ and 5′-GGAATTAGATCTCTAC- CAGC-3′); and GFP–LGALS8-R232H (5′-CATGAGATCTACCGCCACCA-3′ and 5′-GGAATTAGATCTCTACCAAC-3′) were obtained by gene synthesis (In- tegrated DNA Technologies) based on Ensembl transcript ENST00000526589. The LGALS8 constructs were obtained through the same cloning strategy as per- formed for GFP–PLA2G16.

Immunoblotting 3 In short, cells were lysed in sample buffer, separated by SDS–PAGE and transferred onto polyvinylidene fluoride (PVDF) membranes (Millipore) by wet western blotting. Membranes were blocked with 4% non-fat milk powder (Merck) in PBS 0.1% Tween-20 (Biorad) and subsequently incubated with a primary PLA2G16 antibody (1:500), and ATG7 (1:500) to confirm the absence of endogenous protein. Antibodies against CDK4 (1:1,000) and Rictor (1:1,000) were used as a loading control. After washing, membranes were incubated with secondary antibodies coupled to HRP followed by analysis on a gel imaging system (Bio-Rad). Cleavage of GFP from GFP–LGALS8 was detected with a GFP antibody. For the eIF4G cleavage assay cells (wild-type and ∆PLA2G16) were seeded 1 day before infection. The next day, cells were infected (MOI of ∼10) for 2 h with PV1 or CV-B1, respectively (either with or without pre-incubation for 30 min with 2 mM GHL). Cells were lysed in sample buffer and western blot analyses were subsequently performed using antibodies against eIF4G (1:1,000) and α-tubulin (1:1,000).

Immunofluorescence microscopy For all infection experiments, cells were seeded on IbiTreat μ-slide 18-wells (Ibidi). The following day, virus was bound on ice for 1 h, after which the cells were washed with ice-cold PBS, and subsequently refreshed with complete medium. Next, cells were incubated at 37 °C for indicated time points to let virus internalize into the cells. Cells were then fixed with PBS containing 4% paraformaldehyde and further processed. All cells were counterstained with Hoechst33342. For detection of

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VP1, or dsRNA, cells were permeabilized with 0.3% Triton X-100 in PBS, blocked with blocking buffer (1% BSA, 10% goat serum, 0.3% Triton X-100 in PBS) and stained with the primary antibody J2 anti-dsRNA (1:500)36, or VP1 (1:50) for 1 h at room temperature. Cells were then washed three times with PBS and stained with secondary antibodies coupled to Alexa Fluor 488 for 30 min and counterstained with Hoechst33342. Lysosomal disruption after LLOme (200 μM) treatment was visualized with a LAMP1 antibody (1:1,000) and subsequently stained with the secondary antibody coupled to Alexa Fluor 568. Cy5 labelling of PV1 was adapted from ref. 37. In short, sucrose gradient purified PV1 was incubated with an amine-reactive dye (GE Healthcare) in PBS for 2 h at room temperature. Next, unbound dye was removed using gel filtration columns (Nap5; GE Healthcare), after which virus titres were determined with plaque assays. Infections were performed in the presence or absence of pleconaril (1 μM, Sigma Aldrich) In situ hybridizations were performed as previously described31. In short, after infection, cells were stained with 48 fluorescent RNA probes complementary to the genome of PV1 (accession number NC_002058.3), or CV-B1 (accession number NC_001472.1) according to manufacturer’s specification (Stellaris RNA FISH, Biosearch Technologies). In case of a co-staining, specific antibody against VP1 (1:50) was added to the first wash step, and cells were incubated with secondary antibodies coupled to Alexa Fluor 488 (Life Technologies) during the second wash step. Live cell experiments were performed on glass bottom Willco Wells in a heated, CO2-regulated chamber. Entry assays were all imaged, and analysed on a LeicaMicrosystems confocal microscope using LCS software, and brightness and contrast were adjusted using Adobe Photoshop.

Transmission electron microscopy PV1 (MOI of ∼10) was bound for 1 h on ice, cells (wild-type, ∆PLA2G16, and reconstituted ∆PLA2G16 cells) were washed and incubatedat37°Cfor6h.Cellswer ethenfixedusingKarnovsky’sfixative(2.0% paraformaldehyde 2.5% glutaraldehyde in 100 mM cacodylate buffer at pH 7.2), followed by post-fixation with 1% osmiumtetroxide in 100 mM cacodylate buffer. Next, samples were washed, stained and blocked with Ultrastain 1 (Leica). Then, cells were subjected to ethanol dehydration series, embedded in DDSA/NMA/ Embed-812 (EMS) and sectioned. Sections were stained with Ultrastain 2 (Leica) and analysed on a CM10 electron microscope (FEI).

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In vivo experiments All animal studies were performed under the supervision and with the approval of our local experimental animal committee at The Netherlands Cancer Institute (DECNKI), and comply with local and international regulations and ethical guidelines. ∆Pla2g16 mice were created with the use of a C57BL/6N embryonic stem (ES) cell line (clone EPD0350_5_A06, Eucomm) that contains a targeting cassette spanning exon 2 to 4. This cassette contains a splice acceptor and polyadenylation sequence, causing the Pla2g16 gene product to yield a truncated protein product. The ES cells were injected into blastocysts as described before38 and chimaeric offspring was bred to generate homozygousPla2g16 tm1a(EUCOMM) offspring. All genotypes (wild-type, heterozygote, and homozygote Pla2g16 knockout mice) were obtained in the expected Mendelian ratios after breeding. Insertion of the cassette was confirmed using primers surrounding the locus, or imbed- 3 ded in the cassette (primer 1: 5′-GACTAGCTTCTTCAGGTTAA-3′, primer 2: 5′-GCCATATCACATCTGTAGAG-3′, primer 3: 5′-CTGTGGTCCTTGA GTCCTTA-3′). To confirm the absence of PLA2G16 protein, 6-week-old mice (wild-type and homozygote Pla2g16 knockout mice) were euthanized and organs were collected and homogenized in ice-cold RIPA buffer (25 mM Tris-HCl, pH 7.6, 150 mM NaCl, 1% NP-40, 1% sodium deoxycholate, 0.1% SDS) supplemented with protease inhibitor cocktail (Roche). Western blot analyses were subsequently performed using antibodies against PLA2G16 (1:500, Abnova) and α-tubulin (1:1,000). To confirm the previously described obesity phenotype, heterozygous ∆Pla2g16 mice were bred to generate (6 weeks old, males only) wild-type and homozygous littermates, which were weighed before the start of the experiment, and subsequently fed a high-fat diet (45% fat; Special Diet Services) for 50 weeks and weighed each week (only wild-type and homozygous Pla2g16 knockout mice were used for the experiment). Next, litters of 1-day-old pups (wild-type and ∆Pla2g16 mice) were injected with CV-A10 (plaque-forming units (p.f.u.) = 3,000), CV-A4 (p.f.u. = 3,000) or PBS only. The experiment was performed blinded. Animals were genotyped after the experiment. Each day after infection, pups (males and females) were examined for paralysis, overall fitness, and morbidity. Pups that were eaten by the mother, or that died 1 day after infection were not included in the experiments. Pups from both genotypes were also euthanized on day 3 after infection for immunohistochemistry. Entire pups were fixed with formalin, dehydrated, embedded in paraffin, and sectioned. Sections were then stained with J2-ds-RNA antibody (1:500).

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Pore-formation assay HeLa cells were seeded in 48-well plates 1 day before the experiment (wild-type, ∆PLA2G16, and reconstituted ∆PLA2G16 cells). The next day, cells were starved for 1 h in medium lacking methionine and cysteine (ThermoFisher Scientific), infected with PV1 on ice for 1 h (MOI of ∼50), washed three times with ice-cold PBS and incubated at 37 °C with fresh pre-warmed medium containing alpha-sarcin toxin (100 μg ml−1)19 for 90 min. Cells were subsequently pulsed with 35S labelled methionine and cysteine (7.4 MBq ml−1) for 20 min. Cells were then washed four times with ice-cold PBS and lysed in Laemmli buffer. Next, lysates were separated using SDS–PAGE electrophoresis. Gels were dried and exposed to a phosphorimager plate BAS-TR2040 and images were taken using a BASReader.

CV-B3 luciferase assay HeLa cells (wild-type, ∆PLA2G16) were seeded in 96-well plates 1 day before infection. Cells were either infected with CV-B3-luc virus for 30 min, or transfected with a CV-B3-luc replicon (15 ng RNA, Lipofectamine2000, ThermoFisher Scientific), after which the medium was replaced with fresh medium (either with/ without preincubation with 2 mM GHL (Sigma Aldrich) for 30 min). Luminescence was measured 6 h after infection or transfection to assess translation of the viral genome. Significance was calculated by a pairedt -test.

Transfection and infection of PV1-DsRed Wild-type, ∆PLA2G16 and reconstituted ∆PLA2G16 HeLa cells were seeded 1 day before infection in 48-well plates. PV1-DsRed RNA was obtained through in vitro transcription of the infectious clone pPVM-2A50-DsRed. In short, DNA was linearized with EcoRI and transcribed with T7 RNA polymerase (MEGAscript T7 Transcription Kit, ThermoFisher Scientific). Next, cells were either infected with PV1-DsRed for 30 min or transfected with PV1-DsRed RNA (30 ng purified RNA was transfected using Lipofetamine2000 (ThermoFisher Scientific)). After 30 min, medium was replaced (either with/without 2 mM GHL). Cells were fixed 10 h after transfection for imaging.

Virus infectivity assay To study the infectivity of picornaviruses in the absence of PLA2G16, LGALS8 and ATG7, HeLa cells were seeded in 48-well plates 1 day before infection. The next day, cells (wild-type, ∆PLA2G16 and reconstituted ∆PLA2G16 cells, and ∆ATG7, ∆PLA2G16/∆ATG7, ∆LGALS8, and ∆PLA2G16/ ∆LGALS8 HeLa cells) were

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infected with an MOI of ∼5. Cells were fixed (4% PFA in PBS) 8 h after infection and stained with J2 anti-dsRNA (1:500), for 1 h at room temperature. Cells were then washed three times with PBS and stained with secondary antibodies coupled to Alexa Fluor 488 for 30 min and counterstained with Hoechst33342. dsRNA- positive cells were assessed using a fluorescence microscope (Zeiss). The mean of the number of dsRNA-positive cells ± s.d. for four fields was calculated using ImageJ, and brightness and contrast were adjusted using Adobe Photoshop. Crystal violet assays were performed as follows. HAP1 (wild type, ∆PLA2G16, reconstituted ∆PLA2G16 cells, and ∆PLA2G16 cells reconstituted with PLA2G16- C113A) and HeLa cells (WT, ∆PLA2G16, ∆ATG7, ∆PLA2G16/∆ATG7, ∆LGALS8, and ∆PLA2G16/∆LGALS8 cells) were seeded in 96-well plates and challenged with the virus of interest at an MOI of ∼5. All viruses were allowed to be absorbed for 1 h, cells were washed with PBS and replication was allowed to occur for 36 h. Cells were subsequently washed with PBS, fixed (4% PFA in PBS) and stained with 3 crystal violet (0.5% crystal violet, 25% methanol in H2O).

Hypotonic shock Cells were treated for 10 min with hypotonic medium (0.5 M sucrose in PBS, with 10% PEG1000) to induce sterile damage to endocytic vesicles. Next, cells were washed twice with PBS and incubated with 60% H2O in PBS for 3 min. Cells were allowed to recover for 20 min in complete medium at 37 °C before fixation and imaging, or cells were lysed at indicated time points for analysis with western blot. The same protocol was followed for live cell imaging, with the exception that the acquisition started during the recovery period.

Data availability statement All unprocessed sequence files of the screens in this paper have been made available at the NCBI Sequence Read Archive (SRA) under the accession number SRP092541. All other data are available from the corresponding author upon reasonable request.

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mechanism of guanidine e ect on the 2C function and evidence for the importance of 2C oligomerization. J. Mol. Biol. 236, 1310–1323 (1994). 21. Maier, O., Marvin, S.A., Wodrich, H., Campbell, E.M. & Wietho, C.M. Spatiotemporal dynamics of adenovirus membrane rupture and endosomal escape. J. Virol. 86, 10821–10828 (2012). 22. Gao, W., Ding, W. X., Stolz, D. B. & Yin, X. M. Induction of macroautophagy by exogenously introduced calcium. Autophagy 4, 754–761 (2008). 23. Hosokawa, N., Hara, Y. & Mizushima, N. Generation of cell lines with tetracycline-regulated autophagy and a role for autophagy in controlling cell size. FEBS Lett. 580, 2623–2629 (2006). 24. Zádori, Z. et al. A viral phospholipase A2 is required for parvovirus infectivity. Dev. Cell 1, 291– 302 (2001). 25. Dorsch, S. et al. The VP1 unique region of and its constituent phospholipase A2-like activity. J. Virol. 76, 2014–2018 (2002). 26. Hughes, P. J. & Stanway, G. The 2A proteins of three diverse picornaviruses are related to each other and to the H-rev107 family of proteins involved in the control of cell proliferation. J. Gen. Virol. 81, 201–207 (2000). 3 27. Teterina, N.L., Levenson, E.A.&Ehrenfeld, E. Viable polioviruses that encode 2A proteins with uorescent protein tags. J. Virol. 84, 1477–1488 (2010). 28. Wessels, E., Duijsings, D., Notebaart, R.A., Melchers, W.J.G. & vanKuppeveld, F. J. M. A proline- rich region in the coxsackievirus 3A protein is required for the protein to inhibit endoplasmic reticulum-to-golgi transport. J. Virol. 79, 5163–5173 (2005). 29. Lanke, K. H. W. et al. GBF1, a guanine nucleotide exchange factor for Arf, is crucial for coxsackievirus B3 RNA replication. J. Virol. 83, 11940–11949 (2009). 30. Zoll,J. et al. Saffold virus, a human Theiler’s-like cardiovirus, is ubiquitous and causes infection early in life. PLoS Pathog. 5, e1000416 (2009). 31. Blomen, V. A. et al. Gene essentiality and synthetic lethality in haploid human cells. Science 350, 1092–1096 (2015). 32. Langmead, B., Trapnell, C., Pop, M. & Salzberg, S. L. Ultrafast and memory-efficient alignment of short DNA sequences to the human genome. Genome Biol. 10, R25 (2009). 33. Jae, L. T. et al. Virus entry. Lassa virus entry requires a trigger-induced receptor switch. Science 344, 1506–1510 (2014). 34. Sanjana, N. E. et al. A transcription activator-likee effector toolbox for genome engineering. Nat. Protocols 7, 171–192 (2012). 35. Cong, L. et al. Multiplex genome engineering using CRISPR/Cassystems. Science 339, 819– 823 (2013). 36. Weber, F., Wagner, V., Rasmussen, S. B., Hartmann, R. & Paludan, S. R. Double-stranded RNA is produced by positive-strand RNA viruses and DNA viruses but not in detectable amounts by negative-strand RNA viruses. J. Virol. 80, 5059–5064 (2006). 37. Brandenburg, B. et al. Imaging poliovirus entry in live cells. PLoS Biol. 5, e183 (2007). 38. Pettitt, S. J.et al. Agouti C57BL/6N embryonic stem cells for mouse genetic resources. Nat. Methods 6, 493–495 (2009).

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A cknowledgements

The authors thank T. Sixma, L. Wessels, S. Nijman, G. Superti-Furga, W. Fischl, G. Casari and members of the Brummelkamp laboratory for discussions and reading of the manuscript; R. Bin Ali for assistance with generating knockout mice; and K. Kirkegaard and H. Ploegh for providing reagents. This work was supported by SNSF Fellowship PA00P3_145411 to E.V.C., and funding from the Cancer Genomics Center (CGC.nl), Nederlandse Organisatie voor Wetenschappelijk Onderzoek (NWO)–VIDI grant 91711316, European Research Council (ERC) Starting Grant (ERC-2012-StG 309634) to T.R.B. Work in the lab of F.J.M.V.K. was supported by NWO-VICI grant 91812628.

Author contributions J.S., F.J.M.V.K., A.P., J.E.C. and T.R.B. were responsible for the overall design of the study, J.S. and J.E.C. carried out the host factor screens, V.A.B. performed the bioinformatics analysis, L.G.V.D.H. assisted in mouse experiments and the generation and characterization of knockout cell lines, M.B. performed the suppressor screen, J.N. contributed to microscopy analysis, H.J. performed electron microscopy studies, E.V.C. analysed in vitro experiments, J.B. and H.J.T. helped with preparation of virus stocks, luciferase assays and preparation of the EV-A71 screen. J.S. and T.R.B. wrote the manuscript, all authors commented on the manuscript.

Author information Reprints and permissions information is available at www.nature.com/reprints. The authors declare competing financial interests: details are available in the online version of the paper. Readers are welcome to comment on the online version of the paper. Correspondence and requests for materials should be addressed to T.R.B. ([email protected]). Reviewer Information Nature thanks J. Bergelson, S. Lemon and the other anonymous reviewer(s) for their contribution to the peer review of this work.

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E xtended data figure 1. Identification and validation ofLA P 2G16 as a picornavirus host factor. a, A haploid genetic screen to identify critical host factors for CV-B3 infection. b, Disruptive (sense orientation, blue) and undisruptive (antisense orientation, yellow) gene-trap insertions mapped to the genomic locus containing phospholipases PLA2G16, RARRES3 and HRASLS2 in different picornavirus screens. An unselected HAP1 dataset is shown for comparison. Disruptive gene-trap integrations are enriched in the PLA2G16 locus. c, Wild-type HAP1 cells, PLA2G16 gene-trapped cells, and PLA2G16 gene-trapped cells expressing cDNA of either wild-type PLA2G16 or PLA2G16- C113A were infected with an increasing amount of PV1 and CV-B1. Surviving cells were stained using crystal violet (n=3). d, Western blot analysis for the presence of PLA2G16 in different cell lines. CDK4 and Rictor were used as a loading control. For gel source data, see Supplementary Fig. 1. e, Cell lines deficient forPLA2G16 and the respective cells expressing Flag-tagged mouse Pla2g16 were infected with PV1 and stained with a dsRNA antibody to determine infectivity. Cells transduced with a control lentiCRISPR lacking a gene-specific gRNA served as control (values denote mean ± s.d., n = 3). f, Wild-type HeLa cells, ∆PLA2G16 cells, and ∆PLA2G16 cells expressing PLA2G16 cDNA, were infected with different picornaviruses to determine dependency on PLA2G16. Cells were stained 8 h after infection with a dsRNA antibody, and infection efficiencies were quantified (mean ± s.d., n = 4, chi-square test, ****P < 0.0001).

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E xtended data figure 2. The generation and characterization ofPla2g16 -deficient mice. a, Overview of Pla2g16-targeted locus. Insertion of a trapping cassette, containing a splice acceptor and polyadenylation sequence, perturbs the Pla2g16 gene. b, Western blot analysis for the presence of PLA2G16 in different wild-type and ∆Pla2g16 mouse tissues. α-Tubulin was used as a loading control. For gel source data, see Supplementary Fig. 1. c, Wild-type and homozygous ∆Pla2g16 mice were exposed to a high-fat diet for a period of 50 weeks. d, Weight of wild-type (n = 10) and homozygous ∆Pla2g16 (n = 11) mice on a high-fat diet (error bars represent s.d.). e, Wild-type and ∆Pla2g16 mouse-tail fibroblasts were infected with CV-A10 and stained with a dsRNA antibody to determine infectivity (mean ± s.d., n = 3, chi-square test, ****P < 0.0001).

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E xtended data figure 3. Characterization of PLA2G16 function during viral infection. a, Detection of Cy5-labelled PV1 (MOI of ∼20, red) virus bound to the cell surface of wild-type and ∆PLA2G16 HeLa cells. Cells were co-stained with wheat-germ agglutinate (WGA, green) to label cell membranes and counterstained with Hoechst33342 (blue) for DNA. ∆PVR cells served as negative control. Insets represent magnifications of dashed boxes.b , Internalization of Cy5-PV1 (MOI of ∼20, red) over time. c, Incubation of Cy5-PV1 (red) in the presence of pleconaril (1 μM) on live GFP–PLA2G16 (green)-expressing HeLa cells, 15 minutes after infection. Individual channels for different regions are shown.d , Electron micrograph of PV1 replication in the indicated genotypes. Cells were infected for 6 h with PV1 (MOI of ∼5). Abundant modification of intracellular membranes was observed in wild-type cells or cells reconstituted with PLA2G16 cDNA. e, Quantification of PV1-DsRed infected and transfected cells (mean ± s.d., n = 3, chi-square test, ****P < 0.0001). f, Yield of viral progeny after transfection of PV1-DsRed. TCID50 was determined on wild-type HeLa cells after 1 cycle of replication (mean± s.d. of three biological replicates).

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E xtended data figure 4. Loss of ATG7 or LGALS8 restores virus susceptibility in ∆PLA2G16 cells. a, Absolute numbers of disruptive mutations in PVR and ATG14 genes in DsRed-negative (low) and -positive (high) channels. Mutation index is determined by the relative number of integrations in the high channel divided by the relative number of integrations in the low channel. b, ∆PLA2G16 HeLa cells expressing GFP–LGALS8 were exposed to PV1, or CV-B1 (MOI of ∼20), leading to the formation of GFP–LGALS8-positive foci. Cells were counterstained with Hoechst33342 (blue) for DNA. Insets represent zoom in of dashed boxes. c, Western blot analysis for the expression of PLA2G16 and ATG7 in indicated genotypes. Rictor was used as a loading control. For gel source data, see Supplementary Fig. 1. d, e, LGALS8 deficiency ∆( LGALS8) or ATG7 deficiency ∆( ATG7) restores virus infection in ∆PLA2G16 HeLa cells. f, ∆ATG7 restores virus infection in ∆PLA2G16 HAP1 cells. The indicated genotypes were infected with an increasing amount of CV-B3– GFP and PV1 (stained with a dsRNA antibody to detect infected cells, n = 4).

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E xtended data figure 5. Viral susceptibility of HeLa cells lacking ATG7 and LGALS8. a, b, Experiments presented in Fig. 3c, d (a) and Extended Data Fig. 4d, e (b) were quantified to determine susceptibility to PV1, and CV-B3–GFP in wild-type, ∆LGALS8, ∆ATG7, ∆PLA2G16, ∆PLA2G16/∆ LGALS8, and ∆PLA2G16/∆ATG7 HeLa cells. Cells were infected, stained with antibodies against dsRNA and counted to determine percentage of infection (mean ± s.d., n = 4, chi-square test, ****P < 0.0001). c, Yield of viral progeny of CV-B3–GFP after one round of replication on indicated genotypes. TCID50 was determined on wild type HeLa cells (mean ± s.d., n = 4).

Extended data figure 6. The N-terminal CRD of LGALS8 is required for recognition of picornavirus induced membrane damage. Indicated genotypes were reconstituted with the N-terminal CRD (R69H), or the C-terminal CRD (R232H) GFP–LGALS8 mutants, with wild-type GFP–LGALS8 serving as a control or with the corresponding empty vector. a, b, Cells were assessed for infectivity with CV-B3 (numbers denote mean ± s.d., n = 4) (a), and for the formation of GFP-positive galectin-8 (GFP–LGALS8) foci during PV1 infection (b). Insets represent magnifications of dashed boxes.

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E xtended data figure 7. PLA2G16 and LGALS8 respond to membrane damage. a, GFP–LGALS8 (green) forms foci when cells are exposed to hypotonic shock in both wild- type and ∆PLA2G16 HeLa cells. Insets represent magnifications of dashed boxes. Untreated cells are shown as a negative control. b, Hypotonic shock induces occasional co-localization of GFP– PLA2G16 and mCherry-LGALS8. Live cells were imaged 10 min after hypotonic shock and individual channels for selected regions are shown. Insets represent magnifications of dashed boxes. c, GFP–PLA2G16 forms foci after hypotonic shock in both wild type as well as ∆LGALS8 cells. Untreated GFP–PLA2G16-expressing cells serve as a negative control. Cells were counterstained with Hoechst33342 (blue) for DNA.

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E xtended data figure 8. PLA2G16 and LGALS8 respond to membrane damage independently. a, b, GFP–PLA2G16 (green) (a) and GFP– LGALS8 (green) (b) expressed in the respective knockout cell lines localize on LAMP1-positive lysosomes (red). c, d, mCherry–LGALS8 (red), and GFP–PLA2G16 (green) colocalize after 30 min of treatment with 200 μM l-leucyl-l-leucine methyl ester (LLOme, which is converted to the lysomotropic

membranolytic form (Leu-Leu)n-OMe (n ≥ 3) polymers by a lysosomal thiol protease, dipeptidyl peptidase I). e, f, LLOme treatment of ∆LGALS8 cells expressing GFP–PLA2G16 (e) and ∆PLA2G16 cells expressing GFP–LGALS8 (f). g, Localization of GFP–PLA2G16 (green) and Cy5- PV1 (MOI of ∼10, red) in live ∆LGALS8 HeLa cells 15 min after infection.h , LLOme treatment of ∆PLA2G16 cells expressing full-length and truncated GFP–PLA2G16. Lysosomes were stained for LAMP1 (red) and cells were counterstained with Hoechst33342 (blue) for DNA. Insets represent magnifications of dashed boxes. Individual channels for different regions are shown.

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3

E xtended data figure 9 | Detection of PV1 genome using single-molecule FISH. a, Frequency of LGALS8 foci in each infected cell 30 min after infection with PV1 n( = 192 and n = 191 cells, for wild type and ∆PLA2G16, respectively) or CV-B1 (n = 189 and n = 191 cells, for wild type and ∆PLA2G16, respectively) shown as a box plot (measured over n = 5 independent experiments). To estimate the significance of the difference between genotypes, an unpaired Welch-corrected t-test was performed between the mean values in each of the five independent experiments.b , In situ hybridization of PV1 RNA (red) bound to wild-type or ∆PLA2G16 cells. Cells were stained with WGA (green) to label cell membranes and counterstained using Hoechst33342 (blue) for DNA. ∆PVR cells served as negative control. Insets represent magnifications of dashed boxes.c , In situ hybridization of PV1 genomes (MOI of ∼100) over time. Indicated genotypes were infected with PV1 and co-stained for VP1 (green) and viral genomes (red). Cells were counterstained with Hoechst33342 (blue) for DNA. d, Protein lysates of PV1 infected cells (MOI of ∼10, 2 h after infection) probed for eIF4G to measure 2A cleavage of eIF4G in the indicated genotypes. GHL was added 30 min before infection to inhibit viral genome replication. For gel source data, see Supplementary Fig. 1.

PS_JS_def.indd 75 08-10-18 08:47 76 Exploring hospice care in the Netherlands

isolation is the gift, all the others are a test of your endurance, of how much you really want to do it.

PS_JS_def.indd 76 08-10-18 08:47 hoofdstuktitel 77 4

Enterovirus D68 receptor requirements unveiled by haploid genetics

Authors: Jim Baggen 1*, Hendrik Jan Thibaut1* , Jacqueline Staring 2, Lucas T. Jae 2, Yue Liu 3, Hongbo Guo 1, Jasper J. Slager 1, Jost W. de Bruin 1, Arno L. W. van Vliet 1, Vincent A. Blomen 2, Pieter Overduin 4, Ju Sheng 3, Cornelis A. M. de Haan 1, Erik de Vries 1, Adam Meijer 4, Michael G. Rossmann 3, Thijn R. Brummelkamp2 , and Frank J. M. van Kuppeveld 1

1 Virology Division, Department of Infectious Diseases and Immunology, Faculty of Veterinary Medicine, Utrecht University, 3584 CL, Utrecht, The Netherlands

2 Biochemistry Division, Netherlands Cancer Institute, 1006 BE, Amsterdam, The Netherlands

3 Department of Biological Sciences, Purdue University, West Lafayette, IN 47907, United States

4 Virology Division, Centre for Infectious Diseases Research, Diagnostics and Screening, National Institute for Public Health and the Environment, 3720 BA, Bilthoven, The Netherlands; and eCancer Genomics Centre, 3584 CG, Utrecht, The Netherlands

*Authors contributed equally

PNAS 2016 vol. 113 (no. 5): 1399–1404

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Enterovirus D68 (EV-D68) is an emerging pathogen that can cause severe respiratory disease and is associated with cases of paralysis, especially among children. Heretofore, information on host factor requirements for EV-D68 infection is scarce. Haploid genetic screening is a powerful tool to reveal factors involved in the entry of pathogens. We performed a genome-wide haploid screen with the EV-D68 prototype Fermon strain to obtain a comprehensive overview of cellular factors supporting EV-D68 infection. We identified and confirmed several genes involved in sialic acid (Sia) biosynthesis, transport, and conjugation to be essential for infection. Moreover, by using knockout cell lines and gene reconstitution, we showed that both α2,6- and α2,3-linked Sia can be used as functional cellular EV-D68 receptors. Importantly, the screen did not reveal a specific protein receptor, suggesting that EV-D68 can use multiple redundant sialylated receptors. Upon testing recent clinical strains, we identified strains that showed a similar Sia dependency, whereas others could infect cells lacking surface Sia, indicating they can use an alternative, nonsialylated receptor. Nevertheless, these Sia-independent strains were still able to bind Sia on human erythrocytes, raising the possibility that these viruses can use multiple receptors. Sequence comparison of Sia-dependent and Sia-independent EV-D68 strains showed that many changes occurred near the canyon that might allow alternative 4 receptor binding. Collectively, our findings provide insights into the identity of the EV-D68 receptor and suggest the possible existence of Sia-independent viruses, which are essential for understanding tropism and disease.

The genusEnterovirus of the family Picornaviridae contains many important pathogens for humans and animals. This genus consists of 12 species: four human enterovirus species (EV-A, -B, -C, and -D), five animal enterovirus species, and three human rhinovirus species. The best known human enterovirus is poliovirus (EV-C), the cause of poliomyelitis and acute flaccid paralysis.O ther well-known enteroviruses are the coxsackieviruses (EV-B and EV-C)—which are the main cause of viral meningitis, conjunctivitis, myocarditis, and —and enterovirus A71, which causes hand-foot-and-mouth disease and is also associated with severe neurological disease, causing serious public health concerns in Southeast Asia (1). Another emerging enterovirus that causes growing public health problems is enterovirus D68 (EV-D68, a member of the species EV-D). Unlike most enteroviruses, which are acid-resistant and multiply in the human , EV-D68 is an acid-sensitive enterovirus (2) that replicates in the respiratory tract. EV-D68 was first isolated from children with respiratory infections in California in 1962 (3).

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Significance

Enterovirus D68 (EV-D68) is an emerging pathogen that recently caused a large outbreak of severe respiratory disease in the United States and is associated with cases of paralysis. Little is known about EV-D68 host factor requirements. Here, using a genome-wide knockout approach, we identified several genes in sialic acid (Sia) biology as being essential for infection. We also showed that not only α2,6-linked Sia, which mainly occurs in the upper respiratory tract, but also α2,3- linked Sia, which mainly occurs in the lower respiratory tract, can serve as the receptor. Moreover, we identified recent EV-D68 isolates that can use an alternative, nonsialylated receptor. Our findings are essential to understand tropism and pathogenesis of EV-D68 as well as the potential of using Sia-targeting inhibitors to treat EV-D68 infections.

It was long considered a rare pathogen, but the frequency of detecting EV-D68 during outbreaks of respiratory disease has increased (4, 5) and over the past decades, three clades of EV-D68 (A, B, and C) have emerged and spread worldwide (6, 7). EV- D68 infections mostly cause mild respiratory disease but can also result in severe bronchiolitis or pneumonia, especially among children (4, 5). In 2014, a nationwide EV-D68 outbreak in the United States was associated with severe respiratory disease and a cluster of acute flaccid myelitis and cranial nerve dysfunction in children, implicating EV-D68 as an emerging public health threat (8, 9). Enteroviruses are small, nonenveloped viruses that contain a single-stranded RNA genome of positive polarity. To initiate infection, enteroviruses bind to specific receptors on host cells. To date, most known enterovirus receptors are cell surface proteins, many of which belong to the Ig superfamily or the integrin receptor family (10). A majority of these receptors bind to the “canyon,” a depression on the virion surface, thereby destabilizing virions and initiating uncoating (11). In EV- D68, the canyon is unusually shallow and narrow, possibly excluding use of large protein receptors (12). Both sensitivity of EV-D68 infection to neuraminidase (NA) treatment and hemagglutina- tion assays point to the use of Sia as the receptor (13, 14). However, beside the role of a terminal Sia residue on the receptor, little is known about the type(s) of Sia that can be used by EV-D68 to infect cells, the composition of the underlying glycan, and whether specific sialylated proteins or glycolipids are required for infection.

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Genome-wide genetic screening in human haploid cells is a powerful tool to reveal host factors involved in entry of various pathogens, including viruses (15, 16). In this study, we performed a haploid screen and demonstrate that genes involved in syn- thesis of sialylated glycans are essential for EV-D68 infection. Furthermore, we show that EV-D68 is able to use α2,6-linked as well as α2,3-linked Sia as a cellular receptor, and we provide the first insights into the composition of the underlying sugar back- bone. Finally, we report the identification of recent EV-D68 isolates that can infect Sia-deficient cells, indicating that these strains can use an alternative receptor.

Results

Multiple genes involved in Sia biology determine susceptibility of cells to EV-D68 infection. We performed a haploid genetic screen (17, 18) by infecting mutagenized human HAP1 cells with the EV-D68 prototype strain Fermon CA62-1. The screen identified nine genes involved in Sia biology (Fig. 1 A and B and Fig. S1), seven of which 4

F ig. 1. A haploid genetic screen for EV-D68 identifies genes involved in synthesis of sialylated glycans. (A) An overview of the different steps in synthesis of Sia and sialylated N-linked glycans. Significant hits related to Sia biology are shown in green. Gal, galactose; GlcNAc, N-acetylglucosamine; Man, mannose; Sia, sialic acid. (B) RefSeq gene structures of CMAS and SLC35A1 are shown with the transcrip- tional orientation pointing from left to right. Gene- trap insertions predicted to disrupt gene function (intronic insertions in sense orientation and insertions mapping to exons) are illustrated for HAP1 cells selected with EV-D68 (selected) and a cell population of similar complexity that had not been selected with EV-D68 (unselected). The FDR-correctedP values for the enrichment of disruptive gene-trap mutations in the EV-D68–selected population are indicated (Materials and Methods). For insertion plots of the other hits, see Fig. S1.

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were in the top 10. Among these nine hits are genes involved in the biosynthesis [UDP-GlcNAc-2-epimerase/ManAc kinase (GNE) and N-acetylneuraminic acid synthase (NANS)] and activation [cytidine monophosphate N-acetylneuraminic acid synthetase (CMAS)] of N-acetylneuraminic acid, the predominant form of Sia in humans. Other hits include transporters that transfer the activated sugars CMP- Sia and UDP-galactose [solute carrier family 35 member A1 and A2 (SLC35A1 and

F ig. 2. Both α2,3- and α2,6-linked Sia can support EV-D68 infection. (A) HAP1 cells treated with NA or HAP1 clones having defects in Sia synthesis were challenged with EV-D68 (Fermon), CV- B3, or IAV-GFP at an moi of 10. Infected cells were visualized by immunostaining and fluorescence microscopy. (B) Cells were infected at an moi of 0.1, and the percentage of infected cells was determined by immunostaining and counting. The mean ± SEM of four biological replicates is shown. (C) Yields of infectious virus after a single cycle of virus production. ERAV, equine rhinitis A virus; EV-D70, enterovirus-D70; EV-D94, enterovirus-D94. The mean ± SEM of three biological replicates is shown. In panels B and C, P values compared with WT were calculated by one-way ANOVA of log-transformed data with Bonferroni correction. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001. (D) ST3GAL4/ST6GAL1DKO cells transfected with plasmids containing ST3GAL4 or ST6GAL1 cDNA were exposed to EV-D68, and yields of infectious virus were measured after a single replication cycle. (E) ST6GAL1KO or ST3GAL4/ST6GAL1DKO cells (ST3/ST6DKO) transduced with retrovirus expressing ST3GAL4 cDNA were exposed to EV-D68 and stained with antibody against the viral capsid.

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SLC35A2)] from the cytosol to the Golgi apparatus and four glycosyltransferases responsible for conjugation of N-acetylglucosamine (GlcNAc) to mannose residues in N-linked glycans [mannoside acetylglucosaminyltransferase 5 (MGAT5)], galactose [beta-1,4-galactosyltransferase 1 (B4GALT1)], and Sia, either via α2,3 linkage [ST3 beta-galactoside alpha-2,3-sialyltransferase 4 (ST3GAL4)] or α2,6 linkage [ST6 beta-galactosamide alpha-2,6-sialyltranferase 1 (ST6GAL1)]. Together, these data provide insights into the identity and composition of the EV- D68 receptor, pointing to an important role of α2,6- and α2,3-linked Sia on N-linked glycans in infection. To confirm the results of the genetic screen, we used mutant cell lines lacking surface expression of Sia (SLC35A1KO and CMASKO) or having a defect in formation of α2,3- and/or α2,6-linked Sia (ST3GAL4KO, ST6GAL1KO, and ST3GAL4/ ST6GAL1DKO). The integrity of these mutant cell lines was confirmed by genetic analysis (Fig. S2 A and B), lectin stainings (Fig. S2C), and infection with Sia- dependent and -independent control viruses. Analysis of the number of infected cells showed that SLC35A1KO and CMASKO cells were highly resistant to influenza A virus (IAV), whereas ST3GAL4/ST6GAL1DKO cells were partially resistant (Fig. 2A), consistent with the broad Sia specificity of IAV (19). In contrast, coxsackievirus-B3, which does not require Sia, could infect all cell lines (Fig. 2 A and B), whereas 4 equine rhinitis A virus, a picornavirus that requires α2,3-linked Sia (20), could infect ST6GAL1KO but not ST3GAL4KO cells (Fig. 2C). Upon characterizing EV- D68 Fermon, we observed that infection was inhibited by NA treatment and almost completely blocked in CMASKO, SLC35A1KO, and ST3GAL4/ST6GAL1DKO cells, both at high (Fig. 2A and Fig. S2D) and low multiplicity of infection (moi) (Fig. 2B). Likewise, little, if any, production of progeny virus was observed in these mutant cell lines (Fig. 2C).

EV-D68 can use both α2,6- and α2,3-linked Sia to infect cells. Specificity forα 2,3- or α2,6-linked Sia can greatly affect tissue tropism of respiratory viruses, as the Sia abundance varies between the upper (mainly α2,6-linked) and lower (mainly α2,3-linked) respiratory tract (21, 22). Identification ofST3GAL4 and ST6GAL1 suggested that both α2,6- and α2,3-linked Sia are used for infection. Indeed, ST6GAL1KO cells were less susceptible to EV-D68, whereas ST3GAL4KO cells were equally susceptible as wild-type cells (Fig. 2 A–C), suggesting a preference of EV-D68 for α2,6-linked Sia. However, EV-D68 can also use α2,3-linked Sia as a receptor, as shown by the observation that ST3GAL4/ST6GAL1DKO cells were more resistant to infection than ST6GAL1KO cells (Fig. 2 A–C). Consistently, ST3GAL4/

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ST6GAL1DKO cells could be rendered more susceptible to EV-D68 by transfection of plasmid containing ST3GAL4 cDNA (and also ST6GAL1), but the effect was small due to the low transfection efficiency in HAP1 cells (Fig. 2D). Upon retroviral transduction with ST3GAL4 cDNA, however, high infection efficiency was observed (Fig. 2E). Identification ofST3GAL4 in the genetic screen is likely due to heterogeneous expression of α2,3- and α2,6-linked Sia, which has been described in human airway epithelial cultures but also occurs in cultured cells, including HAP1 (Fig. S2E). Hence, knockout of ST3GAL4 in cells already expressing α2,6-linked Sia at reduced levels likely conferred resistance to infection. In summary, our data show that both α2,6- and α2,3-linked Sia can be used for infection by EV-D68.

Other EV-D members display a similar Sia preference profile We also investigated the Sia dependency of EV-D70 and EV-D94, two other mem­- bers of the EV-D species. EV-D70 causes outbreaks of hemorrhagic conjunctivitis, which are often associated with neurological disorders (23). EV-D94 has been associated with acute flaccid paralysis (24), but information on the clinical relevance of this virus is scarce. EV-D70 was reported to be NA-sensitive (25), whereas any role of Sia in EV-D94 infection is unknown. Like EV-D68 Fermon, EV-D70 and EV-D94 could not replicate in CMASKO and SLC35A1KO cells. Also, ST3GAL4/ST6GAL1DKO cells were less susceptible to infection than ST6GAL1KO cells (although differences for EV-D70 were less pronounced), suggesting that EV-D70 and EV-D94 can use both α2,6- and α2,3-linked Sia as the receptor (Fig. 2C). Altogether, these data suggest conserved Sia use between EV-D members.

Identification of EV-D68 strains that can infect cells in a Sia-independent manner EV-D68 Fermon was isolated more than 50 y ago and genetically differs from currently circulating strains. Therefore, we investigated the Sia dependence of six recent EV-D68 strains—belonging to the three different EV-D68 clades— isolated from patients with respiratory infections in 2009 and 2010 in the Netherlands (5). To minimize the emergence of genetic variants, these isolates were subjected to a limited number of passages in cell culture before they were analyzed for their ability to infect different HAP1 cell lines. Like EV-D68 Fermon, three clinical strains (670, 2042, and 2284) did not cause (CPE) in Sia-deficient cells (Fig. 3A) and were sensitive to NA treatment, as shown by analysis of virus production in a single cycle (Fig. 3B) or multiple cycles of replication (Fig. S3 A and B). These strains, like Fermon, exhibited some virus production in ST6GAL1KO cells, despite

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4

F ig. 3. Identification of EV-D68 strains that can infect cells independently of Sia. (A) HAP1 clones were infected with different EV-D68 strains and stained with crystal violet. B( ) HAP1 cells treated with NA or CMASKO cells were infected with different EV-D68 strains, and yields of infectious virus were measured after a single cycle of replication. Virus input levels (T = 0) are indicated by a dashed line. ND, not detectable. The mean ± SEM of three biological replicates is shown.

the lack of CPE (Fig. S3A). Strikingly, the other three strains (947, 1348, and 742) were able to replicate in Sia-deficient and NA-treated HAP1 cells (Fig. 3 A and B). Sia independence of these strains was not specific for HAP1 cells, as these viruses also efficiently infected NA-treated A549 and HeLa-R19 cells (Fig. CS3 ). In summary, these data demonstrate that several recent EV-D68 strains strongly depend on Sia, whereas other strains can infect cells in a Sia-independent manner, pointing toward the use of a nonsialylated receptor.

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F ig. 4. A map of the EV-D68 surface residues. Surface representation of the complete EV-D68 capsid structure (12) (Left) and a close-up showing the Sia-binding site within the canyon (Right). The black triangle shows an icosahedral asymmetric unit of the EV-D68 Fermon structure. Residues are colored by radial distance (Å) to the virus center. The black dashed lines outline the canyon in the human rhinovirus 14 structure. Variable residues determining Sia (in)dependence of strains within clade A or clade B are highlighted by black and white contours, respectively. Note that residue 270 in VP1 (black and white contour) is variable both within clade A and clade B. Amino acids were numbered according to Liu et al., 2015 (12).

Sia-independent EV-D68 strains retain Sia-binding capacity A recent study showed that, in vitro, Sia-containing trisaccharides can bind to EV- D68, where the floor of the canyon would be in the major group rhinoviruses or polioviruses (26) (Fig. 4). To gain insight into the residues that allow Sia-independent infection and their location with respect to the Sia-binding site, we sequenced the capsid regions of the different EV-D68 strains. Amino acid sequence comparison showed that the genetically related Sia-dependent (670) and -independent (742 and 1348) strains in clade A differed at 10 positions, whereas the Sia-dependent (2042) and -independent (947) strains in clade B differed at seven positions (Fig. 4 and Tables S1 and S2). Although residues that differed between Sia-dependent and -independent strains in clade A show little overlap with those that were altered in

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clade B, some of the changed residues on the viral surface are near the Sia-binding site (26) (Fig. 4). To investigate whether the amino acid substitutions that established an alternative receptor-binding site might have affected the Sia-binding capacity of these strains, we performed hemagglutination experiments with blood from nine different human donors. All EV-D68 isolates agglutinated human erythrocytes, although two Sia-independent strains (1348 and 742) agglutinated erythrocytes from only one donor (Table 1). Pretreatment of erythrocytes with NA prevented hemagglutination by EV-D68 strains but not by echovirus-7, which agglutinates erythrocytes by binding to its protein receptor, decay-accelerating factor (27). Remarkably, blood from three donors was not agglutinated by any of the EV-D68 strains, whereas hemagglutination titers of IAV were similar for all donors, indicating equal Sia expression levels. This variability suggests that EV-D68 does not merely bind any sialylated glycan but has a preference for specific sialylated glycan structures that are differentially expressed between individuals. No clear correlation between EV-D68 hemagglutination and ABO blood groups was observed. In summary, these data indicate that Sia-independent strains have retained their Sia-binding capacity, albeit two strains (742 and 1348) seem to have a reduced affinity for Sia. 4

Discussion

In this study, we provided important insights into the identity/nature of the EV- D68 receptor. Using a genome-wide haploid screen, we identified genes involved in biosynthesis (GNE and NANS), activation (CMAS), transport (SLC35A1), and conjugation of Sia to glycans (ST3GAL4 and ST6GAL1) as factors required for EV- D68 infection. Using knockout cell lines and gene reconstitution, we have shown that EV-D68 can use both α2,6- and α2,3-linked Sia as a receptor to infect cells. This finding extends recent observations that both α2,6- and α2,3-linked Sia-containing trisaccharides can bind to the EV-D68 capsid and initiate virion uncoating in vitro (26). The observation that EV-D68 can use not onlyα 2,6-linked Sia as a receptor but also α2,3-linked Sia, which resides mainly in the lower respiratory tract, may provide an explanation for its ability to cause severe lower respiratory tract infections. Importantly, the screen did not identify a specific protein receptor, suggesting that EV-D68 can use multiple redundant receptors, given these are glycosylated with a suitable sialylated glycan. Our screen also provided insights into the preference of EV-D68 for specific sialylated glycans. Identification of glycosyltransferases

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responsible for conjugation of galactose (B4GALT1) and GlcNAc (MGAT5) pointed to the importance of Sia–galactose–GlcNAc chains, consistent with the substrate specificity of ST3GAL4 and ST6GAL1. Furthermore, the identification of MGAT5, which forms GlcNAc–β1,6-Man linkages, suggested that EV-D68 spe- cifically recognizes N-linked glycans containing a β1,6-linked antenna. It should be noted that Sia–galactose–GlcNAc, although mainly expressed on N-linked glycans, also occurs on O-linked glycans and glycolipids and that we observed that HEK293S cells, which lack complex N-linked glycans (19), are susceptible to EV- D68. Further proof that EV-D68 does not merely bind any sialylated glycan but has a preference for specific glycans stems from our observation that erythrocytes of several donors could be agglutinated by IAV but not by EV-D68. More research is required to explore the glycan spectrum that can be bound by EV-D68. Upon characterizing recent EV-D68 isolates, we identified strains that are able to infect Sia-deficient cells, implying that these viruses can use an alternative entry receptor. Genetic comparison of Sia-dependent and -independent EV-D68 strains within clades A and B revealed little overlap of residues determining Sia independence but pointed toward residues near the Sia-binding site as possible determinants for Sia (in)dependence. A similar scenario was described for Sia-dependent and -independent strains, where only a few amino acid changes in the Sia- binding site could cause a receptor switch to a nonsialylated glycan (28). It remains to be in- vestigated whether a single amino acid substitution or a combination thereof is required for the observed Sia-independent phenotype. Better understanding of the (combinations of) residues that facilitate Sia independence could ultimately allow prediction of receptor requirement based on sequence alignments. It has been shown in vitro that Sia binds to the EV-D68 canyon at a unique site, compared with the glycan-binding sites in other picornaviruses (Fig. S4). In EV- D68, binding of Sia induces virion destabilization and pocket factor release, which is the first step in the uncoating process (26). We identified several EV-D68 strains that can use a nonsialylated receptor while retaining a Sia-binding capacity, albeit with different affinities. This indicates that the formation of an alternative receptor- binding site does not necessarily result in loss of the Sia-binding site and points to the possible existence of a dual receptor mechanism, where either Sia or a nonsialylated receptor can trigger similar structural conformational changes. How- ever, it is unclear whether binding of Sia to the capsid of Sia-independent viruses still results in virion destabilization and pocket factor release as described for EV- D68 Fermon. Furthermore, more research is warranted to determine whether this alternative receptor is a protein or a sugar that lacks a terminal Sia moiety.

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Although it remains to be established whether Sia-independent strains circulate in the human population, the occurrence of strains that use an alternative receptor could have an impact on tissue tropism and pathogenesis of EV-D68. Also, application of the sialidase DAS-181 (Fludase) (29), an investigational drug against influenza virus that was shown to inhibit EV-D68 (30), may be ineffective against Sia-independent EV-D68 strains. Hence, detailed insight into the interactions of EV-D68 with its receptor(s) is required to understand viral pathogenesis and to develop effective antiviral treatment.

Materials and Methods

Cells and viruses Information on viruses and cells used in this study is described in SI Materials and Methods.

Haploid genetic screen with EV-D68 HAP1 cells were gene-trap mutagenized as described previously (31). Following expansion, 108 mutagenized cells were exposed to EV-D68 Fermon (moi 3). After 4 selection, surviving cells were expanded and used for genomic DNA isolation. Insertion sites identified in cells selected with EV-D68 (yielding 414,290 unique

Table 1. Hemagglutination titers (Log2) of EV-D68 strains on human erythrocytes from nine different donors.

Donor 1 2 3 4 5 6 7 8 9 1+ NA 2 + NA

EV-D68 2042 2 2 0 2 4 0 2 0 6 0 0 EV-D68 947 3 2 0 3 4 0 2 0 6 0 0 EV-D68 2284 2 2 0 3 4 0 2 0 5 0 0 EV-D68 670 3 2 0 3 3 0 2 0 7 0 0 EV-D68 742 0 0 0 0 0 0 0 0 3 0 0 EV-D68 1348 1 0 0 0 0 0 0 0 3 0 0 IAV PR8 9 9 9 10 10 9 9 9 10 0 0 IAV WSN 8 8 8 9 8 8 7 8 8 0 0 Echovirus 7 8 7 ND ND ND ND ND ND ND 8 9

ND, not determined.

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gene-trap insertions mapped to genes) and a population of matched control cells of comparable complexity (495,679 unique gene-trap insertions mapped to genes) were aligned to the human genome not filtering for close reads (31). Subsequently, disruptive insertion sites (in sense orientation of the affected gene or mapping to exons) in significantly identified genes were compared in the two cell populations, and P values for enrichment were calculated using a Fisher’s exact test as described previously (31). Disruptive insertion sites in virus-selected and control cells were plotted onto the RefSeq gene bodies for the following transcripts: NM_001497 (B4GALT1), NM_018686 (CMAS), NM_001128227 (GNE), NM_002410 (MGAT5), NM_018946 (NANS), NM_006416 (SLC35A1), NM_005660 (SLC35A2), NM_006278 (ST3GAL4), and NM_173216.2 (ST6GAL1).

Generation of knockout cells ST3GAL4KO and SLC35A1KO HAP1 cells have been described (31). CMASKO cells were obtained from Haplogen GmbH. The CRISPR-Cas9 system was used to generate ST6GAL1KO cells. The entireST6GAL1 locus was excised (Fig. S2A), and subclones were analyzed by genotyping (Table S3). ST3GAL4/ST6GAL1DKO cells were obtained by deleting an exonic region in ST3GAL4 from ST6GAL1KO cells using CRISPR-Cas9.

Infectivity assays Cells were infected with virus for 1 h. After incubation for the indicated period, virus titers were determined by end-point dilution. Crystal violet staining was performed at 3 d postinfection. Where indicated, cells were pretreated with NA from Clostridium perfringens (NEB) or from Arthrobacter ureafaciens (Roche) in serum-free medium for 30 min.

Immunofluorescence assays Paraformaldehyde-fixed cells were stained using rabbit anti-capsid serum against EV-D68 Fermon (produced in house; 1:1,000) or a mouse monoclonal antibody against CV-B3 protein 3A (1:100) (32). For characterization with lectins, cells were stained with fluorescein-labeled Sambucus nigra lectin (Vector Laboratories; 1:1,000) and biotinylated Maackia amurensis lectin I (Vector Laboratories; 1:500). Cells were examined by confocal microscopy (Leica SPE-II) or standard fluorescence microscopy (EVOS FL cell imaging system). The number of nuclei was quantified using ImageJ, and the number of infected cells was quantified visually.

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Isolation and sequencing of EV-D68 strains from clinical specimen Monolayers of tertiary monkey kidney cells (tMKs) or human rhabdomyosarcoma (RD) cells were incubated with 250 μL EV-D68–positive clinical material (mixed nose and throat swabs) derived from patients with influenza-like illness or acute respiratory infection (5) and incubated at 34 °C until CPE was observed. EV-D68 strains 4311000670 (clade A; further referred to as 670) and 4311000742 (clade A; 742) were isolated on tMK cells, whereas strains 4310900947 (clade B; 947), 4310901348 (clade A; 1348), 4310902042 (clade B; 2042), and 4310902284 (clade C; 2284) were isolated on RD cells. Viruses were harvested by one freeze-thawing cycle at –80 °C and clarified by centrifugation. Subsequent passages of virus were done on tMK or RD cell monolayers. The full genome of the virus isolates was sequenced from passage 4 for strains 670 and 742 and from passage 2 of the remaining four strains. All infection experiments described in this study were performed with viruses that had undergone one or two more rounds of passage on RD cells. From these viruses, the 5′UTR and capsid region was sequenced. The latter sequences were used for amino acid comparisons shown in Tables S1 and S2. Detailed information on virus passages, sequences, and GenBank accession numbers is described in SI Materials and Methods. As required by Dutch legislation, surveillance studies have to be registered in the Personal Data Protection Act Register of the Personal Data Protection Commission. 4 The influenza surveillance from which the clinical enterovirus D68 isolates were obtained is registered in this register and no further ethical approval was needed for this virologic study because only anonymized virus isolates were used.

Hemagglutination assays The hemagglutination assay was performed using standard methods. Briefly, twofold dilutions of the virus stocks were incubated with an erythrocyte suspension for 16 h at 4 °C.

A cknowledgments

This work was supported by the Cancer Genomics Centre and EU FP7 Marie Curie Initial Training Network “EUVIRNA” Grant 264286 (to F.J.M.v.K.), the Netherlands Organization for Scientific Research Grants NWO-VICI-91812628 (to F.J.M.v.K.) and NWO-VIDI-91711316 (to T.R.B.), European Research Council ERC Starting Grant ERC-2012-StG 309634 (to T.R.B.), and National Institutes of Health Grant AI011219 (to M.G.R.).

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19. de Vries E, et al. (2012) Influenza A virus entry into cells lacking sialylated N-glycans.Proc Natl Acad Sci USA 109(19):7457–7462. 20. Stevenson RA, Huang J-A, Studdert MJ, Hartley CA (2004) Sialic acid acts as a receptor for equine rhinitis A virus binding and infection. J Gen Virol 85(Pt 9):2535–2543. 21. Shinya K, et al. (2006) Avian flu: Influenza virus receptors in the human airway.Nature 440(7083):435–436. 22. Nicholls JM, Bourne AJ, Chen H, Guan Y, Peiris JSM (2007) Sialic acid receptor detection in the human respiratory tract: Evidence for widespread distribution of potential binding sites for human and avian influenza viruses.Respir Res 8:73. 23. Wright PW, Strauss GH, Langford MP (1992) Acute hemorrhagic conjunctivitis. Am Fam Physician 45(1):173–178. 24. Kabue JP, et al. (2007) New enteroviruses, EV-93 and EV-94, associated with acute flaccid paralysis in the Democratic Republic of the Congo. J Med Virol 79(4):393–400. 25. Alexander DA, Dimock K (2002) Sialic acid functions in enterovirus 70 binding and infection. J Virol 76(22):11265–11272. 26. Liu Y, et al. (2015) Sialic acid-dependent cell entry of human enterovirus D68. Nat Commun 6:8865. 27. Powell RM, et al. (1998) Characterization of echoviruses that bind decay accelerating factor (CD55): Evidence that some haemagglutinating strains use more than one cellular receptor. J Gen Virol 79(Pt 7):1707–1713. 28. Hu L, et al. (2012) Cell attachment protein VP8* of a human rotavirus specifically interacts 4 with A-type histo-blood group antigen. Nature 485(7397):256–259. 29. Malakhov MP, et al. (2006) Sialidase fusion protein as a novel broad-spectrum inhibitor of influenza virus infection.Antimicrob Agents Chemother 50(4):1470–1479. 30. Rhoden E, Zhang M, Nix WA, Oberste MS (2015) In vitro efficacy of antiviral compounds against enterovirus D68. Antimicrob Agents Chemother 59(12):7779–7781. 31. Jae LT, et al. (2013) Deciphering the glycosylome of dystroglycanopathies using haploid screens for lassa virus entry. Science 340(6131):479–483. 32. Dorobantu CM, et al. (2014) Recruitment of PI4KIIIβ to coxsackievirus B3 replication organelles is independent of ACBD3, GBF1, and Arf1. J Virol 88(5):2725–2736. 33. Belov GA, Fogg MH, Ehrenfeld E (2005) Poliovirus proteins induce membrane association of GTPase ADP-ribosylation factor. J Virol 79(11):7207–7216. 34. Lanke KHW, et al. (2009) GBF1, a guanine nucleotide exchange factor for Arf, is crucial for coxsackievirus B3 RNA replication. J Virol 83(22):11940–11949.

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and you’ll do it despite rejection and the worst odds and it will be better than anything else you can imagine.

PS_JS_def.indd 94 08-10-18 08:47 hoofdstuktitel 95 5

KREMEN1 is a host entry receptor for a major group of enteroviruses

Authors: Jacqueline Staring1, Lisa G. van den Hengel1, Matthijs Raaben2, Vincent A. Blomen2, Jan E. Carette3, and Thijn R. Brummelkamp1,4,5.

Affiliations: 1Oncode Institute, Netherlands Cancer Institute, Plesmanlaan 121, 1066 CX Amsterdam, the Netherlands

2Netherlands Cancer Institute, Plesmanlaan 121, 1066 CX Amsterdam, the Netherlands

3Department of Microbiology and Immunology, Stanford University School of Medicine, 299 Campus Drive, Stanford, CA 94305, USA

4CeMM Research Center for Molecular Medicine of the Austrian Academy of Sciences, 1090 Vienna, Austria

5CGC.nl, Netherlands Cancer Institute, Plesmanlaan 121, 1066 CX Amsterdam, the Netherlands

Cell Host & Microbe 2018 23, 1-8

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Summary

Human type A Enteroviruses (EV-As) cause diseases ranging from hand-foot- and-mouth disease to polio-myelitis-like disease. Although cellular receptors are identified for some EV-As, they remain elusive for the majority of EV- As. We identify the cell surface molecule KREMEN1 as an entry receptor for coxsackievirus A10 (CV-A10). Whereas loss of KREMEN1 renders cells resistant to CV-A10 infection, KREMEN1 overexpression enhances CV-A10 binding to the cell surface and increases susceptibility to infection, indicating that KREMEN1 is a rate-limiting factor for CV-A10 infection. Furthermore, the extracellular domain of KREMEN1 binds CV-A10 and functions as a neutralizing agent during infection. Kremen-deficient mice are resistant to CV-A10-induced lethal paralysis, emphasizing the relevance of Kremen for infection in vivo. KREMEN1 is also essential for infection by a phylogenetic and pathogenic related group of EV-As. Collectively these findings highlight the importance of KREMEN1 for these emerging pathogens and its potential as an antiviral therapeutic target.

Introduction

The Enterovirus genus is one of the best-known genera of the Picornavirus family afflicting millions of people each year. Coxsackievirus A10 (CV-A10) belongs to the Enterovirus type A (EV-A) species and has, together with other members such 5 as enterovirus A71 (EV-A71), CV-A16, and CV-A6, been identified as a disease- causing pathogen during recent outbreaks in Asia and Europe (Blomqvist et al., 2010; Lu et al., 2012; Mirand et al., 2012; Yang et al., 2015; Chen et al., 2017). The disease burden of many of these viruses is substantial, with symptoms ranging from rather mild conditions such as herpangina and hand-foot-and-mouth disease (HFMD) to more serious symptoms such as aseptic meningitis (Lu et al., 2012; He et al., 2013; Liu et al., 2014; Yang et al., 2015; Li et al., 2016). Like all members of the picornavirus family, EV-As are small non-enveloped viruses that possess a positive-sense single-strand RNA genome, which needs to be released into the cytosol to initiate infection. To do so, enteroviruses interact with a host-encoded receptor to reach the required environment for uncoating, pore formation, and eventual genome release into the cytosol (Tuthill et al., 2010). Receptor availability often restricts virus infection (Mendelsohn et al., 1989) and

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F igure 1. A haploid genetic screen in human cells reveals KREMEN1 as CV-A10 host factor (A) Table of selected human EV-As and their entry receptor requirements. (B) Schematic overview of the genetic screen to identify host factors for CV-A10 in infected haploid HAP1 cells. (C) Analysis of the genetic screen. Every circle represents an individual gene and the size of the circles corresponds to the number of gene-trap mutations recovered from the virus-selected cell population. KREMEN1 (pink) and PLA2G16 (green) were found to have most insertions compared with an unselected control dataset (1,172 and 2,046, respectively). The y axis represents the significance

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of enrichment of insertions in the affected gene compared with a non-selected control population. Genes are distributed randomly across the x axis. (D) Crystal violet staining of HAP1 wild-type (WT), KREMEN1-null (∆KRM1, TALEN clone), and KREMEN1-null cells expressing cDNA of hKRM1-HA, infected with PV-1, CV-B1, and CV-A10. (E) Crystal violet staining of HeLa wild-type, ∆KRM1 (lenti-CRISPR), and ∆KRM1 cells expressing cDNA of mKRM1-HA, infected with PV-1, CV-B1, and CV-A10 (represents n = 3).

can determine tissue and/or species tropism (Ren et al., 1990). Therefore, virus- receptor interactions are considered to be attractive targets for antiviral therapeutics (Tan et al., 2014). Despite their clinical relevance, the host factors required for infection by this group of viruses are not well studied. In fact, for the majority of this class of enteroviruses the entry receptors are unknown (Figure 1A). Using a haploid genetic screening system in human cells, we here identify KREMEN1 as an essential host factor for CV-A10 infection. KREMEN1 is a cell surface molecule that regulates WNT signaling by binding to DKK and LRP5/6, thereby promoting uptake of this complex through clathrin-mediated endocytosis (Mao et al., 2002). We show that KREMEN1 acts as a limiting factor during CV- A10 infection and that the extracellular domain of KREMEN1 directly binds to CV- A10 virions. We also validate that Kremen is relevant in an in vivo infection model and confirm that it encodes a receptor not just for CV-A10 but also for (at least) six other members of the EV-A species, underlining that KREMEN1 is an important host factor for a major subclass of EV-As. 5 RS ESULT

KREMEN1 is a host factor for CV-A10 infection Previously, we have used insertional mutagenesis in haploid human HAP1 cells to identify host factors for picornaviruses, which pointed out the (mostly known) entry receptors for EV-A71, CV-B1, CV-B3, CV-A7, EV-68, and poliovirus type 1 (PV-1) (Baggen et al., 2016; Staring et al., 2017). To broaden our understanding of the EV-A species, we applied the same strategy to identify genetic host factors for CV-A10 (Figure 1B; Table S1). To do so, we mutagenized a pool of 100 million HAP1 cells using a retroviral gene-trap cassette. This library of mutants was subsequently exposed to CV-A10 to enrich for virus-resistant mutants. To identify enrichment for mutations in genes encoding potential host factors, we

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mapped gene-trap insertion sites in the resistant cell population, assigned them to individual genes, and compared them with a non-selected mutagenized control cell population. As expected, we found PLA2G16 (Staring et al., 2017) to be one of the most frequently mutated genes in the selected population (with 1,172 indepen- dent gene-trap insertion events), as well as additional known picornavirus host factors (PI4KB [Hsu et al., 2010], 116 insertions; C10ORF76 [Blomen et al., 2015], 306 insertions). KREMEN1 was found to be enriched in CV-A10-selected cells with 2,046 independent gene-trap integrations (Figure 1C). This cell surface protein was not identified in the aforementioned screens for picornaviruses, nor was it found as a host factor in similar screens performed in HAP1 cells with viruses representing other viral families (e.g., influenza virus A, adeno-associated virus, and dengue, Ebola, Lassa, Lujo, or respiratory syncytial virus; Carette et al., 2009, 2011; Jae et al., 2014; Marceau et al., 2016; Pillay et al., 2016; Hoffmann et al., 2017; Raaben et al., 2017). To validate the importance of KREMEN1 as a host factor for CV-A10 infection, we generated HAP1 and HeLa cells deficient forKREMEN1 (Figure S1A; ∆KRM1) using TALEN and CRISPR nucleases, respectively. Both ∆KRM1 cell lines were resistant to CV-A10 infection but remained susceptible to other enterovirus serotypes such as PV-1 and CV-B1. Importantly, re-expression of human KRM1-HA (hKRM1-HA) or ectopic expression of mouse Krm1-HA (mKrm1-HA) restored virus susceptibility in these cells, demonstrating that loss of KREMEN1 is solely responsible for the observed resistance phenotype (Figures 1D, 1E, and S1B–S1D). Thus,KREMEN1 is a critical factor for CV-A10 infection in vitro, and its function as host factor is not limited to HAP1 cells.

Kremen null mice are resistant to CV-A10 infection Next, we studied the role of KREMEN1 during viral infection in vivo in mice deficient forKremen1 and Kremen2 (∆Krm1/ ∆Krm2) (Ellwanger et al., 2008). Newborn wild-type and Kremen-deficient mice were infected and examined for typical clinical manifestations of CV-A infection (Gifford and Dalldorf, 1951). As expected, muscle tissue (tongue) of wild-type mice stained positive for double- stranded RNA (dsRNA), indicating virus replication, whereas muscle tissue of ∆Krm1/∆Krm2 animals did not show any sign of virus infection (Figure 2A). Importantly, CV-A10 infection in newborn mice, as with other CV-A viruses, has been shown to lead to viremia, muscle lesions, and (lethal) flaccid paralysis (Gifford and Dalldorf, 1951; Staring et al., 2017). All ∆Krm1/∆Krm2 mice (9 out of 9) that were exposed to CV-A10 remained alive and showed no signs of infection 15 days post-injection, in contrast to the wild-type animals (1 out of 8 injected animals

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F igure 2. Kremen1/2-Deficient Mice Are Resistant to CV-A10 Infection (A) One-day-old wild-type (WT) and Krm1/Krm2 null (∆Krm1/∆Krm2) mice were injected with PBS (uninfected), or injected with CV-A10. Four days after infection, mice were euthanized and muscle tissue (tongue) was stained for double-stranded RNA (dsRNA) to indicate infected (8/1) cells. Scale bar, 100 mm. (B) Wild-type (red line) and ∆Krm1/Krm2 (blue line) WT mice were exposed to CV-A10 and their survival was plotted on a Kaplan-Meier graph. p < 0.001 (log-rank test) for survival.

survived) (Figure 2B). Although this experiment does not distinguish between a role for Kremen1 or Kremen2 in viral infection in vivo, our cell-based experiments did not link KREMEN2, a KREMEN1 homolog described to function redundantly in the WNT signaling pathway (Mao et al., 2002), to virus infection. First, the KREMEN2 locus is, unlike the KREMEN1 locus, not enriched for disruptive mutations in the 5 performed haploid genetic screen (Figure S2A). Furthermore, expression of human KREMEN2-HA (hKRM2-HA) did not restore susceptibility to CV-A10 infection in ∆KRM1 cells (Figures S2B and S2C). Finally, neither expression of mouse Kremen2- HA (mKrm2-HA) nor a fusion construct containing the extracellular domain of mKrm2 attached to the intracellular domain of Krm1m affected CV-A10 binding to the cell surface of ∆KRM1 cells (Figure S2D). This suggests that KREMEN1, and not KREMEN2, is responsible for the resistance to CV-A10 infection.

KREMEN1 expression hypersensitizes cells to CV-A10 When knockout cells reconstituted with a KREMEN1 transgene were exposed to increasing viral titers, we noticed that KREMEN1 overexpression rendered cells more susceptible to viral infection than wild-type cells, suggesting that KREMEN1 is not only required for viral infection, but also can act as a rate-limiting host factor

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F igure 3. KREMEN1 is an entry receptor for CV-A10 (A) Infection assay of wild-type (WT) HeLa cells, ∆KRM1 cells, and ∆KRM1 cells expressing cDNA of mKrm1-HA, infected with an increasing amount of CV-A10. Surviving cells were stained using crystal violet (representation of three experiments). (B) Overexpression of mKrm1-HA makes DKRM1 cells hypersensitive to CV-A10 infection. Wild-type HeLa cells, ∆KRM1, and ∆KRM1 cells expressing cDNA of mKrm1-HA were infected with a different OM I of CV-A10, stained with a dsRNA antibody as a measure for virus replicating cells, and infected cells were quantified (n = 3,

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mean ± SD, chi-square test, ****p < 0.0001). (C) Indicated cell types were infected with CV-A10 for 1 hr on ice and stained for CV-A10 genomes (in situ hybridization, red). Scale bar, 10 mm. (D) Binding of CV-A10 genomes on cell surface of HEK293 and HCT116 cells overexpressing cDNA of mKrm1-HA. mKrm1-HA-positive cells (green) are laced with CV-A10 genomes (red). Scale bar, 10 mm. (E) CV-A10 genomes (red) are detectable in the same compartment as mKrm1- HA (green) 10 min post-infection in the indicated cell types. ∆KRM1 HeLa cells overexpressing mKrm1-HA. Dashed box represents enlarged areas. Nuclei are counterstained with Hoechst (blue). Scale bar, 10 μm. (F) Immunoprecipitation of CV-A10 with empty (mock), LAMP1-Fc-coated, and KRM1-Fc-coated Sepharose beads. CV-A10 were detected with an anti-VP-1 antibody and immunoprecipitated. CV-A10 was used for RT-PCR to detect viral genomes. (G) Competition assay with KRM1-Fc- and LAMP1-Fc-conditioned medium. CV-A10 and CV-B1 viruses were incubated on ice with different concentrations of KRM1-Fc or LAMP1-Fc protein for 1 hr before infection. Cells were stained with dsRNA 7 hr post-infection. Infected cells were quantified and plotted after normalization against infected controls in unconditioned medium (n= 3, mean ± SD).

(Figures 3A and 3B). Because KREMEN1 is a transmembrane protein that resides at the cell surface (Mao et al., 2002), we examined whether KREMEN1 could act as a virus receptor. To study binding of infectious CV-A10 particles to the cell surface, we incubated CV-A10 for 1 hr on ice with ∆KRM1 cells, ∆KRM1 cells expressing mKrm1-HA, and wild-type HeLa cells. Subsequently, these cells were stained for CV-A10 genomes by performing in situ hybridization using specific fluorescent probes. ∆KRM1 cells showed a consistent decrease in viral genome binding compared with wild-type HeLa cells, whereas mKrm1-HA-overexpressing cells showed increased binding of viral particles containing CV-A10 genome (Figures 3C and S2E). Elevated binding of CV-A10 virions to cells overexpressing mKrm1-HA 5 was also observed in other cell lines, such as HEK293 and HCT116 cells (Figure 3D). Subsequent incubation of cells at 37 C led to co-internalization of CV-A10 with mKRM1-HA (Figure 3E), suggesting that KREMEN1 binds to virions at the cell surface and that this complex would subsequently undergo endocytosis.

CV-A10 binds the extracellular domain of KREMEN1 To address whether KREMEN1 could directly bind to CV-A10 virions, we purified the extracellular domain of human KREMEN1 fused to the Fc region of rabbit immunoglobulin G (hKRM1-Fc). Incubation of CV-A10 with immobilized hKRM1-Fc protein indeed resulted in the co-precipitation of virions. Importantly, when the ectodomain of the Lassa virus receptor LAMP1 (Jae et al., 2014) was used as bait, no binding was observed (Figure 3F). To confirm that the KREMEN1/CV- A10 interaction is required for infection, we incubated CV-A10 with an increasing

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F igure 4. KREMEN1 serves as a receptor for multiple CV-A viruses (A–I) KREMEN1 dependency of different CV-A viruses. Wild-type (WT) HeLa cells, ∆KRM1 cells, and ∆KRM1 cells expressing cDNA of mKrm1-HA were infected with CV-A2 (A), CV-A3 (B), CV- A4 (C), CV-A5 (D), CV-A6 (E), CV-A12 (F), CV-A10 (G), CV-A7 (H), and CV-A14 (I). Cells were infected for 7 hr and stained with a dsRNA antibody, after which infection efficiencies were quantified (mean ± SD, n = 4, chi-square test; **p < 0.01; ****p < 0.0001; ns, not significant). (J) Cladogram of the P1 genomic region of serotypes found in the human EV-A species (only serotypes of which the full P1 region is known were used). Bootstrap values (0–1) are shown. Green, KREMEN1 dependency; purple, unknown receptor; red, SCARB2: SCARB2 was linked to all viruses in this group, whereas PSGL-1 and Annexin II were previously linked to EV-A71. Asterisk indicates that the virus was not tested. See also Figure S3.

amount of hKRM1-Fc- or LAMP1-Fc-conditioned medium prior to infection. Indeed, hKRM1-Fc, but not LAMP1-Fc, incubation led to inhibition of infection in a concentration-dependent manner. In line with these findings, hKRM1-Fc did not affect CV-B1 infection (Figure 3G). Thus, CV-A10 virions bind KREMEN1 on cells, leading to subsequent endocytosis of the KREMEN1/CV-A10 complex.

KREMEN1 is an important host factor for a subset of EV-As As illustrated in Figure 1A, the entry receptor(s) are unknown for a substantial group of EV-As. We therefore explored whether additional EV-A strains were dependent on KREMEN1. Infection assays using CV-A2, -A3, -A4, -A5, -A6, and -A12 demonstrated that these viruses indeed critically relied on KREMEN1 as a host factor (Figures 4A–4G), whereas the infectivity of CV-A7 and CV-A14 was 5 unaffected byKREMEN1 perturbation (Figures 4H, 4I, and S3A), in agreement with the previous findings that these viruses rely on other host factors for entry (Nishimura et al., 2009; Yamayoshi et al., 2009, 2012; Tan et al., 2013; Zhang et al., 2017). Interestingly, sequence analysis of the P1 region (containing the structural proteins) of EV-A genomes revealed that the KREMEN1-dependent viruses form a distinct group within the EV-A species (Figure 4J), suggesting that KREMEN1 functions not just as a critical host factor for CV-A10, but also for a large subset of the EV-As.

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Discussion

Here we assign a function to KREMEN1 as an entry receptor for a large group of enteroviruses, including most of the coxsackie type A viruses. Although outbreaks used to be dominated by EV-71 and CV-A16, recent epidemics have been more dominated by CV-A10, CV-A6, and other EV-A species. While these viruses are responsible for a lesser disease burden, these EV- As still have a major effect on human health (Lu et al., 2012; Mirand et al., 2012; He et al., 2013; Yip et al., 2013; Li et al., 2016). Furthermore, the high occurrence of recombination between enterovirus viral species (Hu et al., 2011) makes it difficult to predict how these promiscuous subtypes will behave during future outbreaks. The host factors required for infection may constitute valuable targets for the development of antiviral therapeutics. Enteroviruses, like other virus species, have been difficult to classify. The discovery of new serotypes, the high mutation rates, and the aforementioned occurrence of recombination between different serotypes of the Enterovirus species lead to frequent renaming and reclassification (Purdy et al., 2017). Taxo- nomic classification can be based on sequence similarity, but can also, as with the ‘‘major’’ and ‘‘minor’’ classes of rhinoviruses, be based on receptor dependency. In addition, receptor requirement may suggest a common strategy for the develop- ment of antiviral therapeutics against this subclass of picornaviruses. The EV-A species has previously been subdivided into three groups based on sequence similarity and pathogenesis (Yamayoshi et al., 2012). The first group, of which EV- A71 is the main member, shares SCARB2 as an entry receptor (Figure 4J, red box) and is associated with cases of HFMD. The second group (Figure 4J, purple box), contains rare EV-As of which the receptors remain unidentified (Yamayoshi et al., 2012). The identification ofKREMEN1 as an essential entry receptor for CV-A10 and as a critical host factor for at least six additional members of the human EV-A species highlights a distinguishing feature for the third subgroup of EV-As (Figure 4J, green box). It will be interesting to address how receptor use affects the clinical manifestations of these viruses.

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Star★methods

Detailed methods are provided in the online version of this paper and include the following:

• Key resources table • Contact for reagent and resource sharing • Experimental model and subject details – In Vivo Experiments B Cell Culture – Viruses • Method details – Haploid Genetic Screen for CV-A10 Host Factors – Generation of KREMEN1 Knock-Out Cell Lines – Reagents and Antibodies – Constructs and Plasmids – Crystal Violet Assay – Immunoblotting – Virus Infection Assays – In Situ Hybridizations – CV-A10 Interaction with KREMEN1 – Competition Assay – Polygenetic Analysis for Human Type A Enteroviruses • Quantification and statistic ananalysis • Data and software availability 5 – Data Availability Statement

Supplemental information Supplemental information includes three figures and one table and can be found with this article online at https://doi.org/10.1016/j.chom.2018.03.019.

A cknowledgments

The authors thank J. Nieuwenhuis and other members of the Brummelkamp laboratory for discussions and reading of the manuscript, and M. Enders and C. Niehrs for providing valuable reagents. We would also like to thank P. Celie from the protein facility of the Netherlands Cancer Institute for his assistance with the

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production of the recombinant proteins used in this study. This work is part of the Oncode Institute, which is partly financed by the Dutch Cancer Society and was funded by the gravitation program CancerGenomiCs.nl from the Netherlands Organization for Scientific Research (NWO), NWO-VICI grant (016.Vici.170.033), and European Research Council starting grant (ERC-2012-StG 309634) to T.R.B.

Author contributions J.S. and T.R.B. were responsible for the overall design of the study; J.S. and J.E.C. carried out the host factor screen; V.A.B. performed the bioinformatics analysis; L.G.v.d.H. assisted in mouse experiments, the generation and characterization of knockout cell lines, and the cloning of expression constructs; M.R. assisted in the pull-down assays; J.S. and T.R.B. wrote the manuscript; and all authors commented on the manuscript.

Declaration of interests T.R.B. is co-founder and advisory board member of Haplogen, and co-founder and managing director of Scenic Biotech.

Received: December 13, 2017 Revised: February 19, 2018 Accepted: March 26, 2018 Published: April 19, 2018

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Smith, D.B.; ICTV Report Consortium (2017). ICTV virus taxonomy profile: picornaviridae. J. Gen. Virol. 98, 2645–2646. 25. Raaben, M., Jae, L.T., Herbert, A.S., Kuehne, A.I., Stubbs, S.H., Chou, Y., Blomen, V.A., Kirchhausen, T., Dye, J.M., Brummelkamp, T.R., et al. (2017). NRP2 and CD63 are host factors for Lujo virus cell entry. Cell Host Microbe 22, 688–696. 26. Ren, R., Costantini, F., Gorgacz, E.J., Lee, J.J., and Racaniello, V.R. (1990). Transgenic mice expressing a human poliovirus receptor: a new model for poliomyelitis. Cell 63, 353–362. 27. Sanjana, N.E., Cong, L., Zhou, Y., Cunniff, M.M., Feng, G., and Zhang, F. (2011). A transcription activator-like effector (TALE) toolbox for genome engineering. Nat. Protoc. 7, 171–192. 28. Sanjana, N.E., Shalem, O., and Zhang, F. (2014). Improved vectors and genome-wide libraries for CRISPR screening. Nat. Methods 11, 783–784. 29. Staring, J., Von Castelmur, E., Blomen, V.A., Van Den Hengel, L.G., Brockmann, M., Baggen, J., Thibaut, H.J., Nieuwenhuis, J., Janssen, H., van Kuppeveld, F.J.M., et al. (2017). PLA2G16 represents a switch between entry and clearance of Picornaviridae. Nature 541, 412–416. 30. Tan, C.W., Poh, C.L., Sam, I.C., and Chan, Y.F. (2013). Enterovirus 71 uses cell surface heparan sulfate glycosaminoglycan as an attachment receptor. J. Virol. 87, 611–620. 31. Tan, C.W., Lai, J.K.F., Sam, I.C., and Chan, Y.F. (2014). Recent developments in antiviral agents against enterovirus 71 infection. J. Biomed. Sci. 21, 14. 32. Terletskaia-Ladwig, E., Meier, S., Hahn, R., Leinmueller, M., Schneider, F., and Enders, M. (2008). A convenient rapid culture assay for the detection of enteroviruses in clinical samples: comparison with conventional cell culture and RT-PCR. J. Med. Microbiol. 57, 1000–1006. 33. Tuthill, T., Groppelli, E., Hogle, J.M., and Rowlands, D.J. (2010). Picornaviruses. Curr. Top. Microbiol. Immunol. 343, 43–89. 34. Yamayoshi, S., Yamashita, Y., Li, J., Hanagata, N., Minowa, T., Takemura, T., and Koike, S. (2009). Scavenger receptor B2 is a cellular receptor for enterovirus 71. Nat. Med. 15, 798–801. 35. Yamayoshi, S., Iizuka, S., Yamashita, T., Minagawa, H., Mizuta, K., Okamoto, M., Nishimura, 5 H., Sanjoh, K., Katsushima, N., Itagaki, T., et al. (2012). Human SCARB2-dependent infection by coxsackievirus A7, A14, and A16 and enterovirus 71. J. Virol. 86, 5686–5696. 36. Yang, Q., Ding, J., Cao, J., Huang, Q., Hong, C., and Yang, B. (2015). Epidemiological and etiological characteristics of hand, foot, and mouth dis- ease in Wuhan, China from 2012 to 2013: outbreaks of coxsackieviruses A10. J. Med. Virol. 87, 954–960. 37. Yip, C.C.Y., Lau, S.K.P., Woo, P.C.Y., Wong, S.S.Y., Tsang, T.H.F., Lo, J.Y.C., Lam, W.K., Tsang, C.C., Chan, K.H., and Yuen, K.Y. (2013). Recombinant coxsackievirus A2 and deaths of children, Hong Kong, 2012. Emerg. Infect. Dis. 19, 1285–1288. 38. Zhang, X., Shi, J., Ye, X., Ku, Z., Zhang, C., Liu, Q., and Huang, Z. (2017). Coxsackievirus A16 utilizes cell surface heparan sulfate glycosaminoglycans as its attachment receptor. Emerg. Microbes Infect. 6, e65.

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Star★methods

Key resources table

REAGENT or RESOURCE SOURCE IDENTIFIER Antibodies Anti-VP1 (coxsackievirus A10) M. Enders N/A Anti-HA Sigma-Aldrich Cat# H6908 Anti-eIF4G Cell Signaling Technology Cat# 2498 J2 anti-dsRNA English and Scientific Consulting Cat# 10010200; RRID: AB_2651015 Hoechst33342 Thermo Fisher Scientific Cat# H3570 Alexa Fluor 488 Thermo Fisher Scientific Cat #A11008 Bacterial and Virus Strains ElectroMAX DH10B Cells Thermo Fisher Scientific Cat# 18290015 Gene-trap retrovirus (Jae et al., 2013) N/A Coxsackievirus A2 European Virus archive (EVA) Strain Fleetwood Coxsackievirus A3 ATCC Strain Oslon Coxsackievirus A4 National collection of pathogenic Strain H08302 509 viruses (NCPV) Coxsackievirus A5 ATCC Strain Swartz Coxsackievirus A6 ATCC Strain Gdula Coxsackievirus A7 ATCC Strain Parker Coxsackievirus A10 ATCC Strain Kowalik Coxsackievirus A12 ATCC Strain Texas Coxsackievirus A14 ATCC Strain G-14 Enterovirus A71 ATCC Strain HP Poliovirus type 1 ATCC Strain Sabin Coxsackievirus B1 ATCC Strain Conn-5 Chemicals, Peptides, and Recombinant Proteins IMDM with HEPES and L-Gln Lonza Cat# 12-722F DMEM 4.5 g/L Glu and L-Gln Lonza Cat# 12-604F Blasticidin Invivogen Cat# ant-bl-05 Puromycin Invivogen Cat# ant-pr-1 PVDF Immobilon-P Millipore Cat# IPVH00010 Turbofectin 8.0 Origene Cat# TF81005 Lipofectamine 2000 Invitrogen Cat# 11668027 Protein A Sepharose 4B Thermo Fisher Scientific Cat# 101041 cOmplete protease inhibitor cocktail Roche Cat# 138468 Triton X-100 Sigma Aldrich Cat# 93443 BSA (Sigma-Aldrich) N/A N/A

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Continued REAGENT or RESOURCE SOURCE IDENTIFIER Custom made fluorescent RNA probes Stellaris RNA FISH/ Biosearch N/A directed against the CV-A10 genome Technologies (accession number AY421767.1) Formamide Thermo Fisher Scientific (Ambion) Cat #AM9342 Formaldehyde Sigma Aldrich Cat #F8775-500ml Accuprime Taq HiFi Thermo Fisher Cat 12346086 M270 streptavidin-coated Dynabeads Life Technologies Cat #65305 Circligase II Epicentre Cat #CL9021K Deposited Data Coxsackievirus A10 selection screen This study SRA: SRP125607 Unselected control dataset (Staring et al., 2017) SRA: SRX223544 Experimental Models: Cell Lines HAP1 (Carette et al., 2011) N/A H1-HeLa ATCC Cat# CRL-1958 HEK293 ATCC Cat# CRL-3216 HCT116 ATCC Cat# CCL-247 Vero E6 ATCC Cat# CRL-1586 RD cells ATCC Cat# CCL-136 Experimental Models: Organisms/Strains Mouse: C57BL/6J The Jackson Laboratory RRID: IMSR_JAX:000664 Mouse: Kremen1/Kremen2 -/- (Ellwanger et al., 2008) N/A Recombinant DNA pBabe-Puro-hKRM1-HA This Study N/A pBabe-Puro-hKRM2-HA This Study N/A phCMV-hKRM1-Fc This Study N/A phCMV-hLAMP1-Fc (Jae et al., 2014) N/A 5 pBabe-Puro-mKrm1-HA This Study N/A pBabe-Puro-mKrm2-HA This Study N/A pBabe-Puro-mKrm2/Krm1fusion This Study N/A pGT-GFP (Carette et al., 2009) N/A pVSV-G (Carette et al., 2009) N/A pGAG-POL (Carette et al., 2009) N/A pAdVantage Promega Cat# E1711 pX330 (Cong et al., 2013) Addgene Cat# 42230 pLentiCRISPR (Sanjana et al., 2014) Addgene Cat# 52961 Software and Algorithms Prism 7.0b GraphPad Software N/A ImageJ NIH N/A Adobe Photoshop Adobe N/A LCS software Leica-Microsystems N/A

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Contact for reagent and resource sharing Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Thijn Brummelkamp (T.Brummelkamp@ nki.nl).

Experimental model and subject details

In vivo experiments The animal ethics committee at the Netherlands Cancer Institute (DECNKI) approved the in vivo experiment according to the Dutch legislation on animal experiments. The experiment was conducted blinded, and animals housed under standard conditions. Each pup (male and female) was genotyped after the experiment. Male C57BL/6J mice were obtained from Jackson laboratory. Kremen1/2 knockout (∆Krm1/2) mice were kindly provided C. Niehrs (Ellwanger et al., 2008) and bred in-house. Litters of one-day old pups (wild-type and ∆Krm1/2) received single intraperitoneal injections containing CV-A10 (plaque-forming units, p.f.u. = 3,000) or PBS. After infection, pups were evaluated daily for paralysis, physical health, and morbidity. Pups that died 24 h post injection or due to infanticide were excluded from the experiment. The sex of the pups was not determined, due to their young age. For immunohistochemical analysis, pups of 4 days of age were euthanized and subsequently fixed in formalin, dehydrated and embedded in paraffin. Paraffin sections were stained with an antibody against J2-dsRNA (1:500).

Cell culture HAP1 cells were derived from the human (male) myeloid leukemia cell line KBM- 7 as previously described (Carette et al., 2011). These cells were maintained in IMDM (Thermo Fisher Scientific) supplemented with 10% fetal calf serum (FCS), penicillin–streptomycin and L-glutamine (Thermo Fisher Scientific) at 37 C. H1- HeLa (female), HEK293 (female) and HCT116 (male), Vero cells (female), and RD cells (female) (all ATCC) were cultured in DMEM (Thermo Fisher Scientific) supplemented with 10% fetal calf serum (FCS), penicillin–streptomycin and L-glutamine (Thermo Fisher Scientific) at 37 C. All cell lines were tested to be negative for mycoplasma. Cell lines were authenticated by genotyping, or obtained from authentic stocks (ATCC).

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Viruses The following type A Coxsackieviruses (CV-A) were used in the study; CV-A2, (strain Fleetwood, European Virus archive (EVA)), CV-A3 (Oslon, ATCC), CV-A4 (H08302 509, national collection of pathogenic viruses (NCPV)), CV-A5 (Swartz, ATCC), CV-A6 (Gdula, ATCC), CV-A7 (Parker, ATCC), CV-A10 (Kowalik, ATCC), CV-A12 (Texas, ATCC), and CV-A14 (G-14, ATCC). Furthermore, enterovirus A71 (EV-A71) (HP, ATCC), poliovirus-1 (PV1) (Sabin, ATCC), and coxsackievirus B1 (CV-B1) (Conn-5, ATCC) were also used. All viruses were expanded on Vero or RD cells and standard plaque assays were performed to determine virus titer.

Method details

Haploid genetic screen for CV-A10 host factors HAP1 cells were mutagenized using a retrovirus containing a gene-trap cassette as described previously (Blomen et al., 2015). In short, 12 T175 flasks were seeded with HEK293 cells and transfected with gene-trap and packaging plasmids (Carette et al., 2009). Retrovirus was harvested 48 h post transfection twice a day for 3 days and concentrated by spinning at 60,000g at 4 C. Forty million HAP1 cells were infected with the gene-trap virus (with 8 μg ml 1 protamine sulfate). Subsequently, the infected HAP1 cells were expanded for 5 days, after which the mutagenized cells were frozen down for later use. The CV-A10 screen was performed by infecting 100 million HAP1 cells with CV-A10 (MOI of ~5) in 14 T175 flasks. Genomic DNA was isolated of 5 the resistant cells after approximately 10 days. The isolated genomic DNA was used to amplify the gene-trap insertion sites using a LAM-PCR protocol as previously described (Blomen et al., 2015). To summarize, a portion of the gene-trap cassettes and a part of the flanking genomic region were amplified with a single biotinylated primer for 120 cycles (8 LAM-PCR’s in total), after which the single-stranded DNA products were captured with streptavidin-coated magnetic beads (M-270, Thermo Fisher Scientific). The beads were then washed and the DNA was ligated to a single stranded DNA linker using Circligase II (Epicentre) as previously described, followed by a final exponential amplification of 18 cycles (Blomen et al., 2015). Products of the final PCR were purified (Qiagen, PCR purification kit) and sequenced on a HiSeq2000 (Illumina) using a custom sequencing primer. Insertion site mapping was performed as previously described (Staring et al., 2017). Briefly, the reads obtained from sequencing were aligned to the human genome (hg19) using bowtie (Langmead

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et al., 2009). Only reads that aligned with zero or a single mismatch were selected as insertion sites. These reads were assigned to non-overlapping regions of protein- coding genes (Refseq hg19) using intersectBed and the unique insertion sites more than 2 base pairs apart were counted. Finally, the number of disruptive integrations (in the transcriptional orientation of the gene) was compared to an unselected HAP1 control dataset previously published (Jae et al., 2013) and available at the SRA (SRP018361: SRA: SRX223544) using a one-sided Fisher’s exact test and P-values were false discovery rate (FDR)-corrected (Benjamini-Hochberg). The results of the experiments are displayed in bubble plots with genes randomly distributed over the x-axis and with the log(significance) on the y-axis.

Generation of KREMEN1 knock-out cell lines We used the gene-editing tools TALENs and clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9 in order to generate single KREMEN1 knock- out HAP1 and HeLa clones, respectively. TALEN pairs were designed to target exon 2 in the human KREMEN1 gene and subsequently assembled following the TALE construction protocol (Sanjana et al., 2011) using the TALE toolbox kit (Addgene). TALENs were then transfected into HAP1 cells using Turbofectin (Origene). Next, cells with integrated TALENs were obtained by Blasticidin (Invivogen) selection. To obtain KREMEN1 knock-out HeLa cells, we designed a guide RNA (gRNA) targeting exon 5 (5’-GACGCTGTTGCATTCGGTAC-3’) and cloned this gRNA into the lentiCRISPR V2Blast (Addgene, no 83480). HeLa cells were subsequently transfected with this lentiCRISPR plasmid using Lipofectamine 2000 (Invitrogen) followed by Blasticidin selection. KREMEN1 deficiency in both cell lines was confirmed by Sanger sequencing (Figure S1). HeLa cells were additionally tested for mRNA levels with primers designed against the gRNA targeted region of the KRM1 gene: Fw 5’-TACGGGGAGGCAGCCAGTAC-3’, Rv 5’-CGCTGACACTGAGGTTGGCC-3’.

Reagents and antibodies Protamine sulfate was obtained from Sigma-Aldrich. Antibodies directed against CV- A10 VP1 (mouse) were a kind gift from M. Enders (Terletskaia-Ladwig et al., 2008). Anti-HA antibody (rabbit) was obtained from Sigma-Aldrich, and eIF4G antibodies were from Cell Signaling Technology. J2 anti-dsRNA antibody was obtained from English and Scientific Consulting. Hoechst33342, Alexa Fluor 488-conjugated secondary antibodies were obtained from Thermo Fisher Scientific. The secondary antibodies conjugated to horseradish peroxidase (HRP) were obtained from Biorad.

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Constructs and plasmids Human and mouse KRM1-HA, KRM2-HA, and KRM1-Fc were generated by gene synthesis (IDT) based on Ensembl transcripts; ENSMUST00000020662.14 for mKrm1, ENSMUST00000046525.8 for mKrm2, and ENST00000400335.8 for hKRM1. KRM1 and KRM2 were digested with EcoRI and SalI and cloned into retroviral vector pBabe-Puro (digested with EcoRI and SalI). hKRM1-Fc was digested and cloned into phCMV using BglII and BamHI. LAMP1-Fc was cloned into phCMV, as described previously (Jae et al., 2014). pCS-mFLAG-Krm2 was a kind gift from C. Niehrs (Mao et al., 2002). TheKRM2 m /KRM1fusion construct was obtained by amplifying the extracellular domain (ECD) of mKrm2 using the following primers: Fw 5’-aagatcgaattcATGGGAACACCTCATCTGCA-3’ and Rv 5’- tagatcgggccccAGAGGCCTGAGCGGCGCCAG-3’ and digesting ECD- mKRM2-HA with EcoRI and ApaI after which the fragment was ligated in pBabe- Puro mKrm1-HA (digested with EcoRI and ApaI).

Crystal violet assay HAP1 (wild-type, ∆KRM1, ∆KRM1+hKRM1-HA) and HeLa cells (wild-type, ∆KRM1, ∆KRM1+mKrm1-HA) were seeded in 48-well plate to be confluent the next day. HAP1 cells were incubated with CV-A10, PV1 or CV-B1 (MOI of ~5) and HeLa cells were incubated with CV-A10, PV1 or CV-B1 (MOI of ~5), or with CV-A10 with increasing MOI, after which virus infection was allowed to proceed at 37 C. After 24 h, viable cells were washed, fixed with 4% PFA/PBS and stained with crystal violet (0.5% crystal violet, 25% methanol in H2O). 5 Immunoblotting In order to detect the presence of KRM1-HA protein, total cell lysates were prepared by lysing HAP1 and HeLa cells (∆KRM1 and ∆KRM1+KRM1-HA) in 6x sample buffer. Proteins were separated by SDS-PAGE and transferred onto polyvinylidene fluoride (PVDF) membranes (Millipore). Membranes were blocked using 5% skim milk powder (Merck) in PBS/0,1% Tween-20 (Biorad) and blotted with an antibody against HA (1:500). EIF4G antibody (1:1000) was used as loading control.

Virus infection assays Staining of viral dsRNA was performed to visualize virus replication in HeLa, and HAP1 cells (wild-type, ∆KRM1, ∆KRM1+mKrm1-HA) after infection with CV- A2, CV-A3, CV-A4, CV-A5, CV-A6, CV-A7, CV-A10, CV-A12, CV14. In short, cells were incubated with virus for 1 h on ice, were washed in PBS, and incubated

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at 37 C to allow replication of virus. After 7 h, cells were fixed with 4% PFA/PBS, washed with PBS and subsequently incubated in blocking buffer (3% BSA/PBS). After blocking, cells were permeabilized using PBS containing 0,3% Triton X-100 (Sigma-Aldrich), 1% BSA (Sigma-Aldrich) and 10% goat serum (Sigma-Aldrich). Next, cells were incubated with the primary antibody J2 dsRNA (1:500) for 30 min at RT. Cells were washed with PBS three times before incubation with Alexa Fluor 488 conjugated secondary antibody. Nuclear staining was subsequently performed with Hoechst33342. DsRNA-positive cells were visualized under a fluorescence microscope (Zeiss) and the average number of dsRNA-positive cells ± SD of 3 or 4 fields was calculated using ImageJ. Images were adjusted in brightness and contrast using Adobe Photoshop.

In situ hybridizations Detection of CV-A10 genomes by in situ hybridizations was performed as described previously (Staring et al., 2017). In brief, HeLa cells were seeded on IbiTreat m-slide 18-wells (Ibidi) and incubated overnight. Cells were inoculated with CV-A10 for 1 h on ice after which the cells were fixed with 4% PFA followed by permeabilization with 70% ethanol (in one instance viruses were allowed to be absorbed for 10 min before fixation). Next, cells were stained with 48 fluorescent RNA probes directed against the CV-A10 genome (accession number AY421767.1) according to manufacturer’s specification (Stellaris RNA FISH, Biosearch Technologies). A specific antibody against HA (1:50, rabbit, Sigma), was added to the first wash for 30 min after which the cells were incubated with secondary antibodies coupled to Alexa Fluor 488 (1:500, Life Technologies) during the second wash step. Cells were imaged on a Leica Micro-systems confocal microscope using LCS software, and fluorescent intensity was determined with ImageJ.

CV-A10 interaction with KREMEN1 Two 15-cm dishes of HEK293 cells were transfected with 30 μg phCMV-hKRM1- Fc or phCMV-LAMP1-Fc in DMEM containing 2% serum using Turbofectin (Origene). 48 h after transfection, medium was harvested, filtered (0.45 μm, Millipore), and incubated with 200 ml slurry of protein A Sepharose beads (Thermo Fisher) for 3 h at 4 C. Beads were centrifuged and subsequently washed three times in PBS with 0.05% Tween-20 and a final wash in PBS, after which the beads were resuspended in 800 ml PBS. Next, 400 ml of slurry was spun down and incubated with a 5.03107 p.f.u. of CV-A10 in NETN buffer (total volume of 500 ml) containing protease inhibitor (Roche) for 2 h at 4 C. Beads were washed in

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NETN buffer (4 times) and beads were divided over 2 tubes; 1) for immunoblot- ting, and 2) for RT-PCR analysis. HRP-conjugated secondary antibody was used to detect the presence of Fc-tagged proteins, and a specific antibody directed against CV-A10 was used to detect virus capsids. The RT-PCR was performed in order to amplify the viral genomic RNA of the bound CV-A10 with the following primers targeting the 3D region: (Fw 5’-GAGGAGGCTGTGTCACTCAT-3’ and Rv 5’ACAGTCGATGGGAAATGGGT-3’;).

Competition assay 15-cm dishes of HEK293 cells were transfected with 30 μg phCMV-hKRM1-Fc or phCMV-LAMP1-Fc in DMEM containing 2% serum using Turbofectin (Origene). 48 h after transfection, medium was harvested, filtered (0.45μ m, Millipore), and the conditioned medium containing the Fc-tagged proteins was run over a column with 50 ml protein A Sepharose beads (Thermo Fisher Scientific). The beads were washed 2 times with 500 ml PBS and eluted with 100 ml 100 mM Glycine buffer (pH 2.7) and collected in 10 ml 1M Tris (pH 8.8) to neutralize the elution buffer. Next, protein samples were run on a SDS-PAGE and protein concentrations were determined by comparison to a standard curve of GST protein samples using a Biorad imaging system. Next, different concentrations of hKRM1-Fc and LAMP1- Fc were added to complete medium as indicated and incubated with a 4,0X105 plaque-forming units (p.f.u.) CV-A10 for 1 h on ice, after which the mixture was used for inoculating wild-type HeLa cells in a 96-wells format for 6 h. Cells were fixed with 4% PFA in PBS and stained with a dsRNA (J2) antibody. Infected cells were visualized with a fluorescence microscope (Zeiss). The mean of the number of 5 dsRNA-positive cells ± SD for 3 fields was calculated using ImageJ, and brightness and contrast were adjusted using Adobe Photoshop.

Polygenetic analysis for human type a enteroviruses The protein sequence of the P1 region (VP4, VP2, VP3, and VP1) was used for the alignment of type A Enteroviruses (that are known to infect humans) for the construction of the cladogram (http://phylogeny.lirmm.fr/phylo_cgi/index. cgi) (Dereeper et al., 2008). CV-A2 (accession number (ACNO) AY421760.1) , CV-A3 (ACNO AY421761.1), CV-A4 (ACNO AY421762.1), CV-A5 (ACNO AY421763.1) CV-A6 (ANCO AY421764.1), CV-A7 (ANCO AY421765.1), CV- A8 (ANCO AY421766.1), CV-A10 (ANCO AY421767.1 ), CV-A12 (ANCO AY421768.1), CV-A14 (ANCO AY421769.1), CV-A16 (ANCO KM055004.1), EV-A71 (ANCO U22522.1), EV-A76 (ANCO JF905564.1), EV-A89 (ANCO

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JX538041.1), EV-A90 (ANCO JX390655.1), EV-A91 (ANCO AY697461.1), EV-A92 (ANCO EF667344.1), EV-A114 (ANCO KU355877.1), and EV-A121 (ANCO NC_030454.1). Other EV-A serotypes were not used for alignment due to limited, or no sequence availability.

Quantification and statistical analysis

Infection assays are expressed as means ± standard deviation (SD). For the comparison of continuous variables from independent groups Chi-square test. All values were normally distributed. The statistical details (n-numbers, mean ± SD and tests) are given in the figure legends and a Log-rank test was used to calculate the P-value of the animal survival experiments. Calculations were performed with Prism version 7.0b software (GraphPad) and Image J.

Data and software availability

Data availability statement All unprocessed sequence files of the screens in this paper have been made available at the NCBI Sequence Read Archive (SRA) under the accession number SRA: SRP125607. All other data are available from the Lead Contact upon reasonable request.

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Supplemental information figure 1. Generation and validation of KREMEN1 knockout cell lines. Related to figure 1 (A) Disrupted alleles of KREMEN1 in HAP1 and HeLa cells using TALEN and CRISPR techniques, respectively. (B) Western Blot analysis for hKRM1-HA expression in HAP1 KRM1-deficient cells. (C) Western Blot analysis for Krm1m -HA expression in HeLa KRM1-deficient cells. (D) RT-PCR forKRM1 using cDNA isolated from WT and KRM1-deficient HeLa cells.

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Supplemental information figure 2. KREMEN2 expression does not compensate for KREMEN1 loss during CV-A10 infection. Related to figure 3. (A) Sense orientation (dark blue, disruptive), and anti-sense orientation (pink, non-disruptive) gene-trap integrations mapped on the KREMEN1 and KREMEN2 loci after CV-A10 infection. Disruptive insertions are only enriched in the KREMEN1 locus. Insertions in KRM1 and KRM2 loci in unselected cells are shown for comparison. (B) HAP1 ΔKRM1 cells expressing hKRM1- HA and hKRM2-HA cDNA were infected with CV-A10 and stained for dsRNA (green) 7 h post infection. Scale bar, 50 μm (C) Indicated cell types expressing mKrm1-HA, or mKrm2-HA were infected with CV-A10 and viable cells were stained with crystal violet. (D) In situ hybridizations of CV-A10 (red) particles bound to HeLa cells expressing mKrm1-HA, mKrm2-HA, or a chimeric Krm2(M1-S339)/Krm1-HA(W377-D473) (mKrm2/Krm1fusion, green). Nuclei (blue) were stained with Hoechst. Scale bar, 10 μm (E) Quantification of CV-A10 genomes bound to WT cells, ∆KRM1 cells, and ∆KRM1 cells expressing mKrm1-HA after 1 h incubation on ice (mean ± SD, two-sided t-test, ** P< 0.005, AU, arbitrary units).

Supplemental Information figure 3. KREMEN1 is required for infection for a subset of CV-A viruses. Related to figure 4. (A) Wild-type HeLa cells, ΔKRM1 cells, and ΔKRM1 cells expressing cDNA of mKrm1-HA were infected with indicated EV-A’s and stained for dsRNA. Nuclei were stained with Hoechst for counter staining (images are representations of 4 replicates). Scale bar, 50 μm. 5

PS_JS_def.indd 123 08-10-18 08:47 124 Exploring hospice care in the Netherlands

if you’re going to try, go all the way. there is no other feeling like that. you will be alone with the gods and the nights will flame with fire.

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General Discussion I: Viral endosomal escape & detection at a glance

Authors: Jacqueline Staring1,4, Matthijs Raaben1, and Thijn R. Brummelkamp1,2,3,4

Affiliations: 1Netherlands Cancer Institute, Plesmanlaan 121, 1066 CX Amsterdam, The Netherlands. 2CeMM Research Center for Molecular Medicine of the Austrian Academy of Sciences, 1090 Vienna, Austria. 3CGC.nl, Netherlands Cancer Institute, Plesmanlaan 121, 1066 CX Amsterdam, The Netherlands. 4Oncode Institute, Netherlands Cancer Institute, Plesmanlaan 121, 1066 CX Amsterdam, The Netherlands.

Journal of Cell Science 2018 vol. 131 (no. 15): jcs216259

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A bstract

In order to replicate, most pathogens need to enter their target cells. Many viruses enter the host cell through an endocytic pathway and hijack endosomes for their journey towards sites of replication. For delivery of their genome to the host cell cytoplasm and to avoid degradation, viruses have to escape this endosomal compartment without host detection. Viruses have developed complex mechanisms to penetrate the endosomal membrane and have evolved to co-opt several host factors to facilitate endosomal escape. Conversely, there is an extensive variety of cellular mechanisms to counteract or impede viral replication. At the level of cell entry, there are cellular defense mechanisms that recognize endosomal membrane damage caused by virus-induced membrane fusion and pore formation, as well as restriction factors that block these processes. In this Cell Science at a Glance article and accompanying poster, we describe the different mechanisms that viruses have evolved to escape the endosomal compartment, as well as the counteracting cellular protection mechanisms. We provide examples for enveloped and non-enveloped viruses, for which we discuss some unique and unexpected cellular responses to virus- entry-induced membrane damage.

Key words

Detection, Endosomes, Escape, Virus

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Introduction

The first step in the life cycle of any virus is the attachment to host cells; for this reason, viruses have evolved to interact with proteins, lipids and sugar moieties on the cell surface, which generally triggers uptake of the virion through the endosomal system (Yamauchi and Helenius, 2013; Cossart and Helenius, 2014). Using this ‘Trojan-horse-like’ entry pathway, viruses gain access to the interior of the target cell, while their genetic material is still shielded from detection by the immune system. By hijacking the endocytic pathways of the host, viruses are able to evade the barrier imposed by the plasma membrane and its connected underlying structures, such as the cortical actin network. Different endocytic cell entry routes for viruses have been described, including clathrin-mediated endocytosis, caveolin- mediated endocytosis, macropinocytosis, and clathrin- and caveolae-independent pathways (Boulant et al., 2015; Matlin et al., 1981; Chardonnet and Dales, 1970; Kartenbeck et al., 1989; Rojek et al., 2008; Nanbo et al., 2010). Endocytosis is highly complex and dynamic, and involves recycling, trafficking, maturation and fusion of endocytic vesicles (Grant and Donaldson, 2009). Viruses that are running the endocytic gauntlet need to make sure they escape the endosomal compartment before being recycled back into the extracellular

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Box 1. Membrane fusion and membrane penetration

Fusion proteins that are embedded in the membrane of enveloped viruses facilitate fusion with cellular membranes, and whereas they can greatly vary in structure, (class I, class II or class III viral fusion proteins) all have a shared mechanism of action: a ligand-triggered conformational change that results in the apposition and eventual merging of the viral lipid envelope and the host membrane (Harrison, 2015). Whereas fusion of two lipid bilayers is thermodynamically favorable, there is, however, a high kinetic barrier (Chernomordik and Kozlov, 2003; Chernomordik et al., 2006). Viral fusion proteins overcome this barrier by utilizing the liberated free energy during the conformational change to pull the viral and host membranes in close proximity through the insertion of fusion loops or peptides, thereby favoring fusion (Bullough et al., 1994; Chen et al., 1999). The different steps that lead to membrane fusion are quite well understood, mainly owing to the availability of crystal structures of pre- and post-fusion conformations of fusion proteins, the best example of which is the hemagglutinin protein from influenza A virus (Skehel and Wiley, 2000; Wilson et al., 1981; Chen et al., 1998) (see poster). Non-enveloped viruses must disrupt the endosomal membrane to access the cytoplasm. However, the mechanism of penetration is not completely understood. Similarly to enveloped viruses, membrane penetration of non-enveloped viruses is driven by conformational rearrangements of viral structural proteins. Three major structural classes of membranolytic viral factors can be distinguished: amphipathic α-helical domains (e.g. adenovirus protein VI; Wiethoff et al., 2005), myristoylated residues (e.g. N-myristoylated VP4 of poliovirus and rhinoviruses; Panjwani et al., 2014; Danthi et al., 2003; Moscufo et al., 1993) and lipid-remodeling enzymatic domains [e.g. parvovirus VP1, which encodes an N-terminal phospholipase type 2 (PLA2); Farr et al., 2005]. Depending on the viral content that needs to be delivered into the cytoplasm (i.e. naked genome or a nucleocapsid), endosomal disruption can be mechanistically accomplished through transient modification of the limiting membrane, formation of a membrane pore, or complete disruption of the endosomal membrane (see poster).

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Box 2. Endovesicular acidity

Acidification has been described as a membrane fusion trigger for orthomyxo­ viruses, rhabdoviruses, α-viruses, flaviviruses, bunyaviruses and (Ochiai et al., 1995; Roberts et al., 1999; Sourisseau et al., 2007; Mosso et al., 2008; Lozach et al., 2010; Cosset et al., 2009), but it is also a well-known trigger for non-enveloped viruses (Bayer et al., 1998; Chung et al., 2005). The vacuolar ATPase (v-ATPase) is a multisubunit complex and proton pump that functions to acidify intracellular compartments (Nelson and Taiz, 1989) (see poster). Its role in virus entry has been well established, and v-ATPase inhibitors such as bafilomycin A1 (Yoshimori et al., 1991) are frequently used as a method of inhibition of cell entry by enveloped viruses that depend on a low pH for membrane fusion. For enveloped viruses, activation of many class I fusion proteins by low pH involves the removal or refolding of globular head domains, which keep the fusion subunit in its pre-fusion state (Godley et al., 1992; Kemble et al., 1992; Rachakonda et al., 2007). Several protonation events are thought to drive this separation; histidine residues of viral capsid proteins, which can function as pH sensors, have especially been implicated here (Kampmann et al., 2006). In addition to membrane fusion proteins, other structural viral proteins facilitate infection of enveloped viruses in a low pH environment. For example, endosomal acidification is not only important for inducing conformational changes in the hemagglutinin fusion protein of influenza A virus, but it also opens up its M2 proton channel, which acidifies the interior of the viral particle (Schnell and Chou, 2008; Stouffer et al., 2008; Cady et al., 2009) (see poster). Subsequently, this dissociates the viral genome (i.e. existing as ribonucleoprotein complexes) from the viral M1 protein, thereby facilitating release from the virion into the host cell cytoplasm (Helenius, 1992). 6

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space (Zila et al., 2014; Mainou and Dermody, 2012; Grove and Marsh, 2011) or subjected to degradation in the harsh environment of the lysosome (see poster). To this end, enveloped viruses (e.g. , Arenaviridae, ) make use of sophisticated molecular machinery that drives fusion of the with the membrane of an endosome, thereby releasing their genomic content into the cytoplasm (Box 1). Non-enveloped viruses on the other hand (e.g. , , Picornaviridae), have evolved different strategies that involve membrane-modifying proteins which can physically pierce the endosomal membrane to allow release of the genomic content into the cytoplasm (see poster). Here, we depict the arms race between viruses and host cells at the level of the endo-lysosome. We highlight the endosomal escape strategies of different viruses and describe the cellular cues that are involved. In addition, we discuss host detection mechanisms at the endo-lysosomal level.

Cues and facilitating factors for endosomal escape A variety of cellular factors trigger the conformational rearrangements in viral capsid proteins that drive membrane fusion and pore formation (see poster). Well- known cues that induce these structural changes include receptor binding, low pH (Box 2), proteolytic processing by endosomal proteases or a combination of these triggers (White and Whittaker, 2016). Moreover, factors also exist to facilitate the entry process. For instance, a switch from cell membrane attachment receptor to intracellular receptor has been described as a process for endosomal escape used by some hemorraghic fever viruses (Jae et al., 2013; Carette et al., 2011). In addition, a specific lipid and/or sugar composition of the endovesicular compartment can be a requirement for viruses to initiate their entry process. However, viruses can often use alternative receptors or entry routes and may respond to multiple proteolytic triggers and entry cues. Key examples for some prototype viruses are described in more detail below.

Receptor binding Conformational changes induced by receptor binding have a big impact on the uncoating and endovesicular escape of several non-enveloped viruses. For example, poliovirus initiates infection by binding to target cells via an interaction with its cellular receptor, the poliovirus receptor (PVR, also known as CD155) (Racaniello, 1996; Mendelsohn et al., 1989). Very early after endocytosis, conformational changes in the poliovirus capsid are promoted by PVR binding (Tsang et al., 2001) and result in exposure of the hydrophobic domains of the capsid protein

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VP1 (Fricks and Hogle, 1990) and the release of the myristoylated VP4 peptide (Chow et al., 1987; Danthi et al., 2003). These conformational changes result in a more hydrophobic capsid, known as the 135S-particle or A-particle (Fenwick and Cooper, 1962; de Sena and Mandel, 1977). This facilitates interaction with the membrane and induces the formation of protein-based transmembrane pores, through which the genomic RNA is believed to be extruded from the early endosome into the cytoplasm (Bostina et al., 2011; Brandenburg et al., 2007; Strauss et al., 2013; Tosteson et al., 2004; Butan et al., 2014). Likewise, adenovirus virions undergo structural rearrangements after binding their receptors [the Coxsackie and adenovirus receptor, CAR (Bergelson et al., 1997) and integrins (Wickham et al., 1993)] and the subsequent endocytosis is thought to mediate complete endosome rupture through the liberation and membrane insertion of viral protein VI in early endosomes (Leopold et al., 1998; Prchla et al., 1995; Martinez et al., 2013; Maier et al., 2012; Meier et al., 2002; Luisoni et al., 2015). For some enveloped viruses (e.g. most Retroviridae, and ), interaction with their cell surface receptor is sufficient to trigger membrane fusion. This reflects their capacity to establish fusion at the plasma membrane at neutral pH, giving them the ability to form enlarged, multi-nucleated cells (syncytia) (Stein et al., 1987; Horvath et al., 1992; Paterson et al., 1985; Morgan et al., 1968). Most enveloped viruses, however, undergo receptor-mediated endocytosis and require a low pH for membrane fusion (see poster).

Proteolytic processing by endosomal proteases A low endosomal pH is also required for Ebola virus cell entry. However, it is not sufficient to trigger membrane fusion (Bale et al., 2011; Brecher et al., 2012); instead, membrane fusion mediated by the Ebola virus glycoprotein involves the digestion by endosomal proteases cathepsin L and cathepsin B (Brecher et al., 2012; Hood et al., 2010; Martinez et al., 2010; Kaletsky et al., 2007; Chandran et al., 2005). There are also non-enveloped viruses that require endosomal proteolysis for cell 6 entry. For example, during reovirus cell entry, after binding of integrin (Maginnis et al., 2006) and JAM-A (Barton et al., 2001) at the cell surface, the capsid protein σ3 is processed and removed by endosomal cathepsins (Baer et al., 1999; Ebert et al., 2002; Doyle et al., 2012), resulting in full exposure of the myristoylated capsid protein μ1, which harbors the membrane penetration activity (Chandran et al., 2002, 2003; Odegard et al., 2004; Nibert et al., 2005). This leads to eventual viral escape from late endosomes (Mainou and Dermody, 2012).

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Intracellular receptor switching Some hemorrhagic fever viruses require additional interactions with endosomal protein receptors in order to escape the vesicular compartment. In the case of Ebola virus, it has been demonstrated that the proteolytic cleavage by endosomal cathepsins, which remove the mucin-like domain and glycan cap of glycoprotein subunit 1, do not fully activate the fusion subunit of the glycoprotein (Bale et al., 2011; Chandran et al., 2005). In addition, after endosomal proteolytic processing, full membrane fusion by Ebola virus glycoprotein occurs in the lysosome and requires a switch from receptors encountered at the cell surface [e.g. DC-SIGN (Alvarez et al., 2002), TIM1 (Kondratowicz et al., 2011)] to an interaction with the intracellular lysosomal membrane receptor Niemann-Pick C1 (NPC1) (Carette et al., 2011; Cote et al., 2011; Wang et al., 2016). Ebola virus membrane fusion also depends on the activity of the homotypic fusion and protein sorting (HOPS) tethering complex, which may orchestrate the correct localization of one or more viral and/or host fusion factors (Jae and Brummelkamp, 2015; Carette et al., 2011). Similarly to Ebola virus, Lassa virus glycoprotein undergoes a conformational change that is induced by low pH, resulting in a receptor switch from α-dystroglycan to lysosome-associated membrane protein 1 (LAMP1) (Jae et al., 2013, 2014; Li et al., 2016). More specifically, this interaction involves the glycoprotein-1 subunit of Lassa virus glycoprotein and an N-glycosylated residue in the distal luminal domain of LAMP1 (Cohen-Dvashi et al., 2015, 2016; Jae et al., 2014). It has recently been suggested that the interaction between the Lassa virus glycoprotein and LAMP1 is specifically responsible for elevating the pH threshold for membrane fusion (Hulseberg et al., 2018). However, for both Lassa and Ebola virus glycoprotein, it remains to be determined whether the interaction with an intracellular receptor is the sole trigger of membrane fusion under physiological conditions, and whether other viruses employ a similar mechanism. Another , Lujo virus, has recently been shown to depend on the endosomal tetraspanin CD63 for membrane fusion (Raaben et al., 2017). Although an interaction with Lujo virus glycoprotein was not reported, it suggests that receptor switching could be a more widespread mechanism for enveloped viruses, and perhaps non-enveloped viruses, to escape the vesicular compartment.

Composition of the endosomal membrane In addition to the cues described above, some viruses have specific lipid or ionic requirements that can influence the efficiency of endosomal escape (Zaitseva et al., 2010). In the case of enveloped viruses, these requirements generally work directly at

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the level of membrane fusion. For example, infection with Semliki Forest virus (SFV) demands an optimal amount of cholesterol and sphingomyelin in the endosomal membrane for fusion (Phalen and Kielian, 1991; White and Helenius, 1980; Nieva et al., 1994). Without these factors, the stable insertion of the fusion loop (which resides within the SFV E1 class II fusion protein) into the target cell membrane is not efficiently established (Ahn et al., 2002). Similarly, as well as a low pH, hantaviruses require high cholesterol levels in the endosomal target membrane for membrane fusion (Kleinfelter et al., 2015). This is probably due to a requirement for the change in fluidity or curvature of the endosomal membrane (Needham and Nunn, 1990; Papanikolaou et al., 2005). Lysobisphosphatidic acid (LBPA), a lipid species which is mostly found in late endosomes and multivesicular bodies (Piper and Katzmann, 2007), is required by several viruses to enter the cytosol. As with cholesterol, the anionic charge and membrane fluidic properties of LBPA appear to be important to facilitate membrane modifications by both enveloped and non-enveloped viruses. For example, the pore-forming activity of the bluetongue virus () Vp5 capsid protein depends on endosomal LBPA levels (Patel et al., 2016) (see poster). Furthermore, zwitterionic phospholipids such as phosphatidylethanolamine and phosphatidylcholine facilitate the structural transition of intermediate subviral particles (from ISVP to ISVP*) of reoviruses (Snyder and Danthi, 2016). Adenovirus has been shown to use a multistep process involving ceramide lipids for robust endosomal membrane penetration (Burckhardt et al., 2011; Meier et al., 2002; Greber et al., 1993, 1996; Wiethoff et al., 2005; Wodrich et al., 2010). After receptor binding, the adenovirus initiates endocytic uptake by generating small lesions in the plasma membrane through release of the viral membranolytic protein VI. These lesions induce a local influx of 2+Ca ions, initiating the membrane-damage repair response. This response entails lysosomal exocytosis and release of acid sphingomyelinase, which degrades sphingomyelin to ceramide lipids in the plasma membrane and stimulates rapid endocytosis of adenovirus particles. Moreover, the ceramide-rich endosomes display enhanced binding of protein VI to the limiting 6 membrane, inducing massive endosomal membrane rupture and subsequent release of the virions (Luisoni et al., 2015) (see poster).

Enzymatic modifications of the endosomal membrane Viruses have also evolved strategies to modify the endosomal membrane through either virus- or host-encoded enzymatic activities to facilitate efficient escape. For example, members of the non-enveloped parvovirus family encode a capsid-tethered lipid-modifying enzyme to alter the endosomal membrane after internalization.

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After receptor binding (e.g. integrins; Weigel-Kelley et al., 2003) and proteolytic processing (via an autolytic process and/or by endosomal cathepsins; Salganik et al., 2012; Akache et al., 2007), parvovirus virions undergo a conformational shift

in late endosomes that leads to the exposure of a phospholipase A2-like domain (PLA2) in their VP1 protein (Suikkanen et al., 2003; Zádori et al., 2001; Farr et al., 2005). This PLA2 domain is capable of damaging liposomesin vitro (Canaan et al., 2004). Furthermore, parvovirus mutants that lack this phospholipase activity have reduced infectivity, but regain their infectivity after incubation with the membrane- permeabilizing agent polyethyleneimine (PEI), or co-incubation with adenovirus virions (Farr et al., 2005). Additional studies report that phospholipase inhibitors prevent parvovirus infection (Dorsch et al., 2002), supporting the current model in which the phospholipase activity in VP1 is indeed required for modification of endosomes and subsequent viral entry into the cytoplasm. In a related fashion, members of the picornavirus family were recently shown to depend on a host- encoded lipid-modifying enzyme for endosomal escape. Picornaviruses, like their prototype poliovirus, recruit cellular phospholipase A2 group XVI (PLA2G16) to virus-induced pores in the endosomal membrane (Staring et al., 2017) (see poster). The catalytic activity of PLA2G16 facilitates the quick release of the viral genome into the cytoplasm, but the exact mechanism by which this is accomplished remains to be determined. Remarkably, the few picornaviruses that are not dependent on PLA2G16 seem to use alternative strategies for membrane penetration, somewhat resembling adenovirus-mediated endosome lysis (i.e. rhinovirus 14; Schober et al., 1998) or encoding a PLA2-like gene (viral 2A protein) in their genome (i.e. parechovirus, avian encephalitis virus and aichivirus; Hughes and Stanway, 2000). Whether these 2A proteins share a similar function as the PLA2 domain of VP1 of parvoviruses remains to be determined. In addition to lipids, it has been suggested that the large carbohydrate network within the endolysosomal lumen, the glycocalyx, also plays a role in viral escape from the vesicular compartment. The neuraminidase protein of influenza A virus strain H5N1 was shown to affect the glycosylation of and LAMP2 at low pH, thereby potentially destabilizing the glycocalyx of the lysosome (Ju et al., 2015).

Cellular antiviral defenses at the level of the endolysomal compartment Viruses are intracellular parasites that depend on their host for multiplication. Whereas the cellular processes described above all favor virus entry and endosomal release of the viral genomic material, there are several systems in play to prevent pathogens from gaining access to the cell interior.

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Protection against incoming virions Antibodies and antiviral peptides, such as defensins, form a first line of defense against incoming virus particles. They can directly interfere with virion binding to receptors and thereby block proper uptake into cells, but also prevent membrane fusion (Leikina et al., 2005) and interfere with capsid destabilization once inside the endosomal compartment (Smith and Nemerow, 2008). For example, α-defensins were shown to bind and stabilize adenovirus particles, subsequently preventing release of protein VI from the capsid and membrane penetration (Smith and Nemerow, 2008). Likewise, θ-defensins inhibit influenza A virus infection by blocking a late step of membrane fusion that is mediated by the viral hemagglutinin protein (Leikina et al., 2005) (see poster). Interestingly, complement component C3 deposited on incoming pathogens co-migrates into endocytic vesicles and can also activate an innate immune signaling cascade (Tam et al., 2014). However, as shown for adenovirus infection, this requires translocation of the capsid–C3 complex into the cytosol. Highlighting the strength of this system, rhinovirus and poliovirus have evolved a mechanism that antagonizes C3 detection by means of proteolytic cleavage of C3 by the virus-encoded 3C protease (Tam et al., 2014). In addition, pattern-recognition receptors can recognize pathogenic factors and form another important line of defense in the endosomal compartment, which is described in more detail elsewhere (Kawai and Akira, 2010; Akira and Takeda, 2004; Lester and Li, 2014).

Detection and inhibition of virus-induced membrane disruption Endosomal membrane modifications that are caused by viral membrane fusion could also directly serve as a danger signal upon which cells respond. Membrane fusion is a process that frequently occurs in cells as part of several physiological processes (e.g. vesicular trafficking). However, virus-induced membrane fusion can be sensed by cells, and this is dependent on the protein stimulator of interferon genes (STING), together with TANK binding kinase 1 (TBK1) and interferon regulatory 6 factor 3 (IRF3) (Holm et al., 2012). Furthermore, influenza A virus has evolved to counteract STING activation through a direct interaction between the fusion peptide of hemagglutinin-2 and STING, highlighting this type of restriction as an important antiviral mechanism (Holm et al., 2016). This implies that cells are able to differentiate virus-induced fusion events from cellular events, or sterile membrane damage. The mechanism(s) by which cells spot viral fusion events as danger signals remains to be established, but this is an intriguing area of innate immune sensing that requires further research. Similarly, members of the interferon-induced

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transmembrane family (IFITM) have been shown to prevent infection of several enveloped viruses at the level of membrane fusion (Desai et al., 2014; Feeley et al., 2011; Li et al., 2013). Three human IFITM proteins (IFITM1-IFITM3) are known to have antiviral activities against several viruses, including dengue virus, Ebola virus, influenza A virus, reovirus, severe acute respiratory syndrome coronavirus (SARS) coronavirus and West Nile virus (Huang et al., 2011; Brass et al., 2009). Mechanistically, there are several lines of evidence indicating that IFITM proteins modify the physical properties of endosomal membranes, thereby inhibiting the formation and/or expansion of fusion pores (Desai et al., 2014; Feeley et al., 2011; Li et al., 2013). Furthermore, overexpression of IFITM3 has been shown to prevent the digestion of reovirus capsid protein μ1, which suggests that IFITM3 can modulate the function of late endosomes, and/or directly interferes with the proteolytic activity that is required for the formation of ISVP* particles (Anafu et al., 2013) (see poster).

Detection and clearance of damaged endosomal membranes Whereas enveloped viruses fuse with endosomal membranes without affecting cell integrity, non-enveloped viruses can cause significant damage to endosomal membranes. This results in exposure of content in the endosomal lumen to cytosolic proteins, thereby presenting a cellular danger signal. For example, endosomal rupture induced by adenovirus protein VI triggers innate pro-inflammatory responses, including activation of the inflammasome (Barlan et al., 2011a,b). In addition, as observed for bacterial infections, the exposure of extracellular carbohydrates such as β-galactoside within the endosomal lumen upon membrane damage can recruit cytosolic galectins. These galectin-positive membranes can be subjected to autophagy for further degradation (Thurston et al., 2012). In the case of adenovirus, galectin-3 and galectin-8 are recruited to sites of protein VI-induced membrane damage, but most adenovirus particles are able to escape the endosomal compartment before being targeted for autophagic degradation (Montespan et al., 2017; Maier et al., 2012) (see poster). Moreover, the removal of damaged membranes in the context of infection could even be beneficial for the virus, because the clearance of the damaged membranes dampens the innate immune response of the infected cell (Zeng and Carlin, 2013). For picornaviruses, it has recently been described that galectin-8 can be recruited to sites of virus-induced membrane damage, which targets these membranes for degradation through the autophagic pathway (Staring et al., 2017). It is currently unclear if the sugar moieties that are recognized by galectin-8 become exposed at the cytoplasmic side, or are detected

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by galectin-8 at the luminal side of the endosome. In order to avoid degradation of their genome, picornaviruses require the activity of the previously mentioned host enzyme PLA2G16. In the absence of this factor, galectin recruitment and subsequent autophagy are sufficient to block infection, through the degradation of the entire virion (including the genome). This additional hurdle in virus entry may very well explain, at least in part, the high particle-to-infectivity ratio of these viruses, where very few viral genomes make it to their expected destination of replication (Bergelson, 2008).

Perspectives Entering the target cell and breaching the host limiting membrane are the first and arguably the most challenging steps for a virus to accomplish. For many viruses, this barrier is represented by the endosomal membrane. An increasing number of host factors that facilitate viral endosomal escape are being identified, and by doing so, mechanistic commonalities get uncovered. For example, the concept of receptor switching seems to be shared by several enveloped viruses (Jae and Brummelkamp, 2015). Future research will determine if intracellular receptor engagement is a common entry strategy for enveloped, and perhaps also non-enveloped, viruses. Another seemingly collective feature of viral endosomal penetration is the ability to co-opt membrane damage responses of the target cell. Whether they involve recruitment of host phospholipases (entry of the picornavirus) or lysosomal exocytosis (entry of the adenovirus), all play a significant role during viral escape from the endosome. Time will tell whether these escape mechanisms are isolated examples, or whether they are widespread and shared by different virus families. Surprisingly little is known about the cellular antiviral response at the level of endosomal membrane penetration, particularly for enveloped viruses. It is well established that cells can sense the viral genomic content once it is released into the cytoplasm through the action of several pattern recognition receptors (Thompson et al., 2011; Jensen and Thomsen, 2012), but are cells able to distinguish between 6 physiological membrane fusion events and virus glycoprotein-triggered membrane fusion? Fundamental questions like these remain to be investigated and it will be interesting to see how our mechanistic knowledge of viral escape and detection pathways continues to expand and provide potential new targets for antiviral strategies.

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Acknowledgements We thank members of the Brummelkamp lab for discussions.

Competing interests T.R.B. is co-founder and shareholder of Haplogen GmbH and Scenic Biotech BV.

Funding This work was supported by funding from the Cancer Genomics Center (CGC.nl), Nederlandse Organisatie voor Wetenschappelijk Onderzoek (NWO)–VICI grant (016.Vici.170.033), European Research Council (ERC) Starting Grant (ERC- 2012- StG 309634) to T.R.B, and by a Marie Sklodowska-Curie Action fellowship (H2020- MSCA-IF-2014 660417) to M.R.

Cell science at a glance A high-resolution version of the poster and individual poster panels are available for downloading at http://jcs.biologists.org/lookup/doi/10.1242/jcs.216259. Supplemental

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75. Li, K., Markosyan, R. M., Zheng, Y. M., Golfetto,O ., Bungart, B., Li, M., Ding, S., He, Y., Liang, C., Lee, J. C. et al. (2013). IFITM proteins restrict viral membrane hemifusion. PLoS Pathog. 9, e1003124. 76. Li, S., Sun, Z., Pryce, R., Parsy, M. L., Fehling, S. K., Schlie, K., Siebert, C. A., Garten, W., Bowden, T. A., Strecker, T. et al. (2016). Acidic pH-induced conformations and LAMP1 binding of the Lassa virus glycoprotein spike. PLoS Pathog. 12, e1005418. 77. Lozach, P.-Y., Mancini, R., Bitto, D., Meier, R.,O estereich, L., Överby, A. K., Pettersson, R. F. and Helenius, A. (2010). Entry of bunyaviruses into mammalian cells. Cell Host Microbe 7, 488-499. 78. Luisoni, S., Suomalainen, M., Boucke, K., Tanner, L. B., Wenk, M. R., Guan, X. L., Grzybek, M., Coskun, and, Greber, U. F. (2015). Co-option of membrane wounding enables virus penetration into cells. Cell Host Microbe 18, 75-85. 79. Maginnis, M. S., , Forrest, J. C., Kopecky-Bromberg, S. A., Dickeson, S. K., Santoro, S. A., Zutter, M. M., Nemerow, G. R., Bergelson, J. M. and Dermody, T. S. (2006). 1 Integrin mediates internalization of mammalian reovirus. J. Virol. 80, 2760-2770. 80. Maier, O., Marvin, S. A., Wodrich, H., Campbell, E. M. and Wiethoff, C. M. (2012). Spatiotemporal dynamics of adenovirus membrane rupture and endosomal escape. J. Virol. 86, 10821-10828. 81. Mainou, B. A. and Dermody, T. S. (2012). Transport to late endosomes is required for efficient reovirus infection. J. Virol. 86, 8346-8358. 82. Martinez, O., Johnson, J., Manicassamy, B., Rong, L., Olinger, G. G., Hensley, L. E. and Basler, C. F. (2010). Zaire Ebola virus entry into human dendritic cells is insensitive to cathepsin L inhibition. Cell. Microbiol. 12, 148-157. 83. Martinez, R., Burrage, A. M., Wiethoff, C. M. and Wodrich, H. (2013). High temporal resolution imaging reveals endosomal membrane penetration and escape of adenoviruses in real time. Methods Mol. Biol. 1064, 211-226. 84. Matlin, K. S., Reggio, H., Helenius, A. and Simons, K. (1981). Infectious entry pathway of influenza virus in a canine kidney cell line. J. Cell Biol.91 , 601-613. 85. Meier, O., Boucke, K., Hammer, S. V., Keller, S., Stidwill, R. P., Hemmi, S. and Greber, U. F. (2002). Adenovirus triggers macropinocytosis and endosomal leakage together with its clathrin-mediated uptake. J. Cell Biol. 158, 1119-1131. 86. Mendelsohn, C. L., Wimmer, E. and Racaniello, V. R. (1989). Cellular receptor for poliovirus: molecular cloning, nucleotide sequence, and expression of a new member of the immunoglobulin superfamily. Cell 56, 855-865. 87. Montespan, C., Marvin, S. A., Austin, S., Burrage, A. M., Roger, B., Rayne, F., Faure, M., Campell, E. M., Schneider, C., Reimer, R. et al. (2017). Multi-layered control of Galectin-8 mediated autophagy during adenovirus cell entry through a conserved PPxY motif in the viral capsid. 13, e1006217. 88. Morgan, C., Rose, H. M. and Mednis, B. (1968). Electron microscopy of . I. Entry. J. Virol. 2, 507-516. 89. Moscufo, N., Yafal, A. G., Rogove, A., Hogle, J. and Chow, M. (1993). A mutation in VP4 defines a new step in the late stages of cell entry by poliovirus.J. Virol. 67, 5075-5078.

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139. Zeng, X. and Carlin, C. R. (2013). Host cell autophagy modulates early stages of adenovirus infections in airway epithelial cells. J. Virol. 87, 2307-2319. 140. 4Zila, V., Difato, F., Klimova, L., Huerfano, S. and Forstova, J. (2014). Involvement of microtubular network and its motors in productive endocytic trafficking of mouse polyomavirus. PLoS ONE 9, e96922.

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do it, do it, do it. do it.

all the way all the way.

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General Discussion II: Concluding remarks

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Identification of novel picornavirus entry receptors Gaining entry into a host cell is a critical and challenging step for a virus. Insights into virus entry and receptor recognition are essential to understand pathogenesis as well as tissue or species tropism. Notwithstanding, the identity of receptors engaged by a variety of viruses is still unknown. In chapter 4 of this thesis we applied a genetic approach to identify host factors important for EV-D68, a virus which can potentially cause severe respiratory illness 1. Several host factors were identified in this screen, including several components of the sialic acid (Sia) biosynthesis pathway, leading to the eventual identification of α2,6- as well as α2,3-linked Sia as entry receptors (as suggested previously 2). Subsequent structural studies indeed demonstrated a direct interaction between EV-D68 and sialylated receptors analogs 3. Furthermore, Sia binding to the canyon of the capsid induced conformational changes and loss of the stabilizing lipid-like moiety or “pocket factor” of the EV-D68 capsid, which is thought to occur in a similar fashion for picornaviruses that interact with immunoglobulin-like receptors 3–5. However, the description of ICAM-5 as a protein receptor for Sia dependent and independent EV-D68 strains 6 could suggests that multiple receptors could be engaged. A similar genetic approach was used in chapter 5, this time to elucidate the host factor requirements for viruses belonging to the Enterovirus type A by selecting cells that were resistant to Coxsackievirus A10 (CV-A10) infection. This screen pointed out KREMEN1, a cell surface receptor that was not implicated in virus infection before. We demonstrate that KREMEN1 can directly bind to CV-A10 virions in vitro, and that it is required for CV-A10 induced pathogenesis in vivo. Surprisingly, KREMEN1 appeared to act as an essential host factor for 6 additional members of the Enterovirus type A family, suggesting that the majority of the Enterovirus type A species use KREMEN1 as an entry receptor. In the absence of KREMEN1 however, HAP-1 cells still displayed residual virus binding at the cell surface of KREMEN1 suggesting attachment to other cell surface molecules. Although binding to these unidentified molecules led to virus internalization, this did not result in successful infection. Unexpectedly, we were unable to demonstrate functionality as an entry receptor for CV-A10 for KREMEN2, a close homologue of KREMEN1 which functions redundantly with KREMEN1 in the WNT signaling pathway 7,8. It will thus be of interest to pinpoint the exact determinants of the interaction between KREMEN1 and CV-A10, and to understand in more detail of how KREMEN1, but not KREMEN2 contributes to the uncoating and subsequent infection of type A 7 Enteroviruses. To conclude, the genetic approaches in haploid human cells can point our new receptor molecules for virus families that are relevant for human health.

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Physiological and pathogenic function of the phospholipase PLA2G16 In chapter 3 we identify PLA2G16 as an important entry factor for picornaviruses. PLA2G16 is a small cytoplasmic phospholipase that in vitro displays various lipid- modifying enzymatic activities including cleavage of the sn-1 or sn-2 positions of glycerophospholipids in addition to an acyltransferase activity. We could demonstrate that its key enzymatic residues were essential for its function as a host factor. Furthermore, using knockout mice we demonstrated that PLA2G16 was important for virus-induced pathology in mice. Using a genetic suppressor screen we unraveled that incoming viruses were subjected to autophagy in PLA2G16-deficient cells through the activation of a pore-sensing pathway that was previously described to clear intracellular bacteria 9. However, although we could show that PLA2G16 localized to incoming virions, and to sites of intracellular membrane damage, the precise molecular role of PLA2G16 during virus infection is not yet understood. In addition, little is known about its function during normal physiological circumstances. Because its presence in mammalian genomes creates a liability for picornavirus infection we suspect that it could have an important physiological role in the absence of viral infection. The high expression levels in adipose tissue in combination with the phenotype of knockout mice have suggested an adipocyte-specific role in lipolysis 10,11. Other studies have implicated PLA2G16 in peroxisome homeostasis 12, where it is seemingly recruited to the peroxisomal membranes 12, but the exact function in adipocytes and its relevance for the biology of peroxisomes remains unclear. Our experiments start to suggest that PLA2G16 may be important during damage occurring at intracellular membranes. Various triggers can cause recruitment of PLA2G16 to organelles other than endosomes. These include damaged lysosomes13 , peroxisomes, mitochondria, as well as structures surrounding the nucleus (unpublished data). Because of our limited understanding of the factors that cause, detect, and repair the damage of membranes it will be interesting to study the function of PLA2G16 in this context. Noteworthily, there are similar lipases belonging to the HRASLS superfamily 14 of which none were scored as host factors in the picornavirus screens presented in this thesis. Furthermore, some picornavirus 2A sequences encode proteins related to the HRASLS family. These viral HRASLS homologs are most similar to PLA2G16 15. This discrepancy between the suggested consanguinity and function of these proteins, may suggest that PLA2G16 has either a higher preference for a specific lipid moiety in comparison to the other lipases, or that it can localize to a specific membrane structure. Until now we have not been able to complement PLA2G16-deficient cells with the viral homologs and it remains unclear if they

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possess any enzymatic activity. However, overexpression of the HRASLS family member RARRES3 did rescue CV-B3 infection in ΔPLA2G16 cells to some extent (unpublished data), suggesting that some of the lipase activity is redundant between the family members. It will be of interest to examine the localization of RARRES3 and other members of the HRASLS family in response to virus infection or membrane damage.

Entry of “non”-enveloped picornaviruses It is striking that the entry pathways of picornaviruses -and poliovirus in particular- remain shrouded in mysteries. While the traditional view entails that non- enveloped viruses exit the cell through a physical disruption of the cell membrane, it has now been shown that PV 16,17, CV-B3 18, and 19 can egress from infected cells through a budding-like process. This new insight will require a revision on how these “enveloped” picornaviruses enter and infect new hosts. Indeed, it was recently demonstrated that cell to cell spreading can occur through these extracellular microvesicles (EMVs) in a more efficient manner compared to single non-membranous virions 17. It also suggests that there might be undiscovered routes involved in their entry. Entry pathways for picornaviruses coated in EMVs (although still dependent on their entry receptor) may require fusion-like events as seen for other membrane coated cargo. One could envision that after such fusion event, entire un-uncoated virions would end up in the cytosol. These virions would not encounter the required “classic” uncoating cues (such as acidification), which are needed to release their genome. This could explain the anti-viral host response through the complement system (as described in chapter 6) 20, and would also suggest that there are other unknown uncoating cues in the cytosol. It could also be that the EMVs lose their integrity at the cell surface thereby allowing the virions to interact with their receptor on the cell membrane. It would be interesting to test if the EMV phenomenon is observed in vivo, if there are different entry cues required for infection, and whether PLA2G16 is required for genome delivery during picornavirus spreading through EMVs. One could speculate that these two versions of spreading (EMV versus naked capsid) may have evolved for different paths of infection; one for the spread between host organisms (as a naked virion which is typically more robust) and one for more efficient spreading between cells in tissues. 7

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Haploid genetics: future perspectives The work presented in this thesis on host factor biology illustrates how loss-of- function genetics can contribute to intriguing new insights in the well-established field of molecular virology. The genome-wide unbiased approach has led to the identification of new picornavirus host factors involved in entry (as show in chapters 3, 4, and 5), but also to the identification of new host factors important for replication (e.g. replication factor C10ORF76 21). Due to the stringency of the method (i.e. the phenotype selected for is cell survival), and due to that fact that many host factors are essential genes (such as ribosomal factors required for translation) that cannot be identified using this system, one arrives at an underappreciation of the host-pathogen network. Because a high intracellular load of viral components also has a negative impact on cell fitness it is challenging (although not impossible22 ) to find host factors that are required for virus egress from the cell. It would thus be beneficial to perform screens with a less harsh selection strategy, as was done with the AAV2-RFP CRISPR screen performed by Marceau and colleagues 23, or by studying only one aspect of the life-cycle, with the use of reporter screens (for instance replicons to study genome replication24). Staining cells with specific antibodies for viral components in combination with FACS sorting might help to elucidate host factors, also for viruses that do not cause cell death. Our studies were performed in HAP1 cells, a cell line that does not necessarily represent any normal tissue, but that has proven to be very useful for expanding our virology knowledge. As hits are in first instance validated using targeted nucleases in HAP1 cells (previously achieved by laborious subcloning of the selected cell population) this also allows for screens performed in knock out backgrounds as was presented in chapter 3, thereby facilitating the study of more complicated genetic interactions by targeting multiple genes in one cell clone. Such an approach could be a suitable route to dissect redundant cellular processes, but as demonstrated in chapter 3, it can also yield unexpected mechanistic insights.

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References

1. Baggen, J. et al. Enterovirus D68 receptor requirements unveiled by haploid genetics. Proc. Natl. Acad. Sci. 113, 1399–1404 (2016). 2. Uncapher, C. R., Dewitt, C. M. & Colonno, R. J. The major and minor group receptor families contain all but one human rhinovirus serotype. Virology 180, 814–817 (1991). 3. Liu, Y. et al. Sialic acid-dependent cell entry of human enterovirus D68. Nat. Commun. 6, (2015). 4. Strauss, M. et al. Nectin-Like Interactions between Poliovirus and Its Receptor Trigger Conformational Changes Associated with Cell Entry. J. Virol. 89, 4143–4157 (2015). 5. Rossmann, M. G. Viral cell recognition and entry. Curr. Sci. 71, 193–204 (1996). 6. Wei, W. et al. ICAM-5/Telencephalin Is a Functional Entry Receptor for Enterovirus D68. Cell Host Microbe 20, 631–641 (2016). 7. Mao, B. et al. Kremen proteins are Dickkopf receptors that regulate Wnt/β-catenin signalling. Nature 417, 664–667 (2002). 8. Staring, J. et al. KREMEN1 Is a Host Entry Receptor for a Major Group of Enteroviruses. Cell Host Microbe 23, 636–643.e5 (2018). 9. Thurston, T. L. M., Wandel, M. P., von Muhlinen, N., Foeglein, A. & Randow, F. Galectin 8 targets damaged vesicles for autophagy to defend cells against bacterial invasion. Nature 482, 414–8 (2012). 10. Duncan, R. E., Sarkadi-Nagy, E., Jaworski, K., Ahmadian, M. & Hei, S. S. Identification and functional characterization of adipose-specific phospholipase A2 (AdPLA).J. Biol. Chem. 283, 25428–25436 (2008). 11. Jaworski, K. et al. AdPLA ablation increases lipolysis and prevents obesity induced by high fat feeding or leptin deficiency.Nat. Med. 15, 159–168 (2010). 12. Uyama, T. et al. Regulation of peroxisomal lipid metabolism by catalytic activity of tumor suppressor H-rev107. J. Biol. Chem. 287, 2706–2718 (2012). 13. Staring, J. et al. PLA2G16 represents a switch between entry and clearance of Picornaviridae. Nature 541, 412–416 (2017). 14. Mardian, E. B., Bradley, R. M. & Duncan, R. E. The HRASLS (PLA/AT) subfamily of enzymes. Journal of Biomedical Science 22, (2015). 15. Hughes, P. J. & Stanway, G. The 2A proteins of three diverse picornaviruses are related to each other and to the H-rev107 family of proteins involved in the control of cell proliferation. J. Gen. Virol. 81, 201–207 (2000). 16. Bird, S. W., Maynard, N. D., Covert, M. W. & Kirkegaard, K. Nonlytic viral spread enhanced by autophagy components. Proc. Natl. Acad. Sci. 111, 13081–13086 (2014). 17. Chen, Y. H. et al. Phosphatidylserine vesicles enable efficient en bloc transmission of enteroviruses. Cell 160, 619–630 (2015). 18. Robinson, S. M. et al. Coxsackievirus B Exits the Host Cell in Shed Microvesicles Displaying 7 Autophagosomal Markers. PLoS Pathog. 10, (2014). 19. Feng, Z. et al. A pathogenic picornavirus acquires an envelope by hijacking cellular membranes. Nature 496, 367–371 (2013).

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20. Tam, J. C. H., Bidgood, S. R., McEwan, W. A. & James, L. C. Intracellular sensing of complement C3 activates cell autonomous immunity. Science 345, 1256070–1256070 (2014). 21. Carette, J. E.et al. Global gene disruption in human cells to assign genes to phenotypes. Nat. Biotechnol. 29, 542–546 (2011). 22. Hoffmann, H.-H.et al. Diverse Viruses Require the Calcium Transporter SPCA1 for Maturation and Spread. Cell Host Microbe 22, 460–470.e5 (2017). 23. Pillay, S. et al. An essential receptor for adeno-associated virus infection. Nature 530, 108–112 (2016). 24. Cherry, S. et al. Genome-wide RNAi screen reveals a specific sensitivity of IRES-containing RNA viruses to host translation inhibition. Genes Dev. 19, 445–452 (2005).

PS_JS_def.indd 158 08-10-18 08:47 PS_JS_def.indd 159 08-10-18 08:47 160 Exploring hospice care in the Netherlands

you will ride life straight to perfect laughter, it’s the only good fight there is.

Charles Bukowski

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Addendum

Summary Nederlandse samenvatting Curriculum vitae List of publications Acknowledgements

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Summary

The family of Picornaviruses is one of the most diverse pathogenic families known to infect humans and animals. As described in chapter 1, these viruses have been studied for more than a century. Surprisingly, many aspects of the viral life cycle still remain unclear, and although for some members of this family vaccination campaigns are highly effective, medications to treat or prevent viral infection are not available. Chapter 2 describes how different genetic techniques can be used to identify host and restriction factors. One of these approaches is mutagenesis-based genetics in haploid human cells.

The strength of this technique is illustrated inchapter 3, where this method is used to unravel the unknown biology of the heavily studied prototype picornavirus; poliovirus. This chapter describes a how multiple phenotypic genetic screens with poliovirus and other enteroviruses lead to the identification of the phospholipase PLA2G16 as a broadly required picornavirus host factor. Chapter 3 continues by applying a genetic suppressor screen in a PLA2G16-null background, which provides insight into why PLA2G16 is recruited to endosomal damage induced by entering picornaviruses. This suppressor screen highlights members of an anti- viral autophagic response pathway, which was previously described for bacterial infections, leading to the hypothesis that PLA2G16 is required for the virus to gain entry into the host before the virus gets sequestered by this previously unknown innate anti-viral immune response.

The potential of haploid genetics is further explored in chapter 4 and chapter 5. Chapter 4 presents a potential new receptor for enterovirus-D68 (EV-D68). In this part of the thesis, the identification of several host factors involved in sialic acid (Sia) biosynthesis leads to the identification of -2,6- and -2,3-linked Sia as receptors for EV-D68. Likewise, chapter 5 presents KREMEN1 as an entry receptor for multiple coxsackie A viruses (CV-A). KREMEN1 is known to be a transmembrane protein involved in the trafficking of Dickkopf through clathrin-mediated endocytosis. In this chapter, data is presented that demonstrates that KREMEN1 can directly interact with CV-A10 as a functional entry receptor. Furthermore, it is shown that KREMEN1 is required for virus infection both in vitro as well as in vivo.

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The above-mentioned experimental chapters highlight the capability of genetic screens to elucidate the importance of host factor biology. Furthermore, these chapters have added to our understanding of the entry pathways used by these viruses and the restriction factors that try to counteract it. Chapter 6 explores the current knowledge of this host-pathogen interplay in a broader context, thereby providing an overview of strategies used by different virus species to escape from the endolysosomal compartment without getting detected. Finally,chapter 7 discusses future perspectives considering the work presented in this thesis.

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N ederlandse samenvatting

De familie van Picornavirussen is één van de meest diverse virusfamilies die zowel mens als dier kan infecteren. In hoofdstuk 1 van dit proefschrift wordt beschreven hoe deze virusfamilie de afgelopen 100 jaar grondig bestudeerd is en wordt er een opsomming gegeven van wat er bekend is van deze virussen. Hieruit blijkt dat verrassend veel aspecten van de levenscyclus van deze virussen tot heden ten dage onbekend zijn. En al zijn er succesvolle vaccinatieprogramma’s opgezet voor een aantal van deze virussen, toch blijft de behandeling van picornavirus infecties met de hulp van medicatie onmogelijk.

Hoofdstuk 2 gaat verder in op de genetische technieken die door de jaren heen gebruikt zijn om picornavirussen te bestuderen. Daarbij wordt de nadruk gelegd op genetica en meer specifiek op de technieken die met de hulp van mutagenese in humane haploïde cellen gastheer factoren kunnen identificeren die nodig zijn voor de infectie van een humane cel.

De kracht van deze techniek wordt onderstreept met de data die gepresenteerd wordt in hoofdstuk 3. In dit eerste experimentele hoofdstuk van dit proefschrift wordt de genetische screenings methodiek toegepast op een aantal picornavirussen, met poliovirus als klassiek voorbeeld voor de familie. Met de hulp van bovengenoemde methodiek wordt een gen geïdentificeerd dat een belangrijke rol speelt tijdens picornavirus infecties in humane cellen. Dit gen en het bijbehorende eiwitproduct draagt de naam PLA2G16. Eerdere studies hebben laten zien dat PLA2G16 fosfolipase-activiteit bezit en dat het een belangrijke rol speelt in de vethuishouding van de muis. Echter, er was nog geen link gelegd tussen dit gen en picornavirus infecties. Hoofdstuk 3 demonstreert dat PLA2G16 niet alleen belangrijk is voor picornavirus infecties in cellen, maar dat verlies van dit gen in de muis ook leidt tot resistentie tegen het virus. Een tweede screen, dit keer in de afwezigheid van PLA2G16, laat vervolgens zien dat picornavirussen het eiwit hoogstwaarschijnlijk nodig hebben om hun genoom snel de gastheercel binnen te loodsen voordat de cel een antivirale immuunreactie activeert middels de inductie van autofagie.

De potentie van haploïde genetica in humane cellen wordt verder bekrachtigd in de experimentele hoofdstukken 4 en 5. In hoofdstuk 4 wordt een enterovirus (een subgroep van picornavirussen) uitgelicht. Voor dit enterovirus D68 (EV-D68), dat gelinkt wordt aan verlammingsverschijnselen bij geïnfecteerde kinderen, was er tot

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nu toe geen cellulaire receptor bekend. Eenzelfde soort genetische aanpak als in hoofdstuk 3 wordt hier toegepast en leidt tot de identificatie van verscheidene genen die betrokken zijn bij Siaalzuur biosynthese, een monosacharide (suikergroep) die op verschillende receptormoleculen op het celmembraan aanwezig zijn. De identificatie van deze genen en de verdere beschreven experimenten suggereren dat deze suikermoleculen kunnen functioneren als receptoren van EV-D68.

In hoofdstuk 5 wordt een andere enterovirus subgroep nader bestudeerd. Deze groep, de type A Enterovirussen (EV-A), veroorzaken voornamelijk hand-, mond-, en voetzweer bij peuters. Ook in dit hoofdstuk leidt het toepassen van de genetische screenings methodiek tot het identificeren van een tot dan toe onbekende receptor. Deze receptor, KREMEN1, blijkt relevant voor 6 leden van de EV-A subfamilie. Hoofdstuk 6 gaat verder in op de directe interactie tussen KREMEN1 en coxsackievirus A10 (CV-A10, lid van de EV-A subfamilie). Ook wordt de in vivo relevantie van KREMEN1 aangetoond met de hulp van infectieproeven in muizen waarbij KREMEN1 afwezig is.

De bovengenoemde experimentele hoofdstukken zijn goede voorbeelden die de kracht van de genetica tonen in context van de virologie. De gebruikte technieken lenen zich uitstekend voor het analyseren en identificeren van de door virussen misbruikte toegangswegen die naar het cytoplasma van de cel leiden. Hoofdstuk 6 schetst een overzicht van de actuele kennis over de interactie tussen een virus en zijn gastheercel. Het hoofdstuk gaat verder in op hoe verschillende virusfamilies ontsnappen uit het endosomale compartiment zonder dat zij door de geïnfecteerde cel gedetecteerd worden. Het laatste hoofdstuk van dit proefschrift,hoofdstuk 7, bediscussieert de experimentele data en gaat in op eventuele conclusies die daaruit voortvloeien.

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Curriculum vitae

Jacqueline Staring werd geboren op 8 december 1984 te Leiden. Na het behalen van haar vwo-diploma is zij in 2004 biomedische wetenschappen gaan studeren aan de universiteit Utrecht. Hier heeft zij in 2007 haar Bachelor of Science behaald. Vervolgens heeft zij op dezelfde universiteit haar studie voorgezet en heeft zij in 2010 haar Master of Science behaald (Cancer Genomics & Developmental Biology). Tijdens deze masteropleiding heeft zij onderzoeksstages afgerond bij prof. dr. M.A.G.G. Vooijs in het Universitair Medisch Centrum Utrecht in Utrecht en bij prof. dr. T.R. Brummelkamp in het Whitehead Institute for Biomedical Research in Cambridge, Massachusetts, in de Verenigde Staten. Uiteindelijk is zij een promotieonderzoek gestart bij prof. dr. Brummelkamp in het Whitehead Institute, hetwelk zij voortgezet heeft op het Nederlands Kanker Instituut te Amsterdam. De resultaten van dit onderzoek worden beschreven in dit proefschrift.

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L ist of publications

Baggen, J., Thibaut, H.J.,Staring, J., Jae, L.T., Liu, Y., Guo, H., Slager, J.J., de Bruin, J.W., van Vliet, A.L.W., Blomen, V.A., et al. (2016). Enterovirus D68 receptor requirements unveiled by haploid genetics. Proc. Natl. Acad. Sci. 113, 1399–1404.

Blomen, V.A., Májek, P., Jae, L.T., Bigenzahn, J.W., Nieuwenhuis, J., Staring, J., Sacco, R., van Diemen, F.R., Olk, N., Stukalov, A., et al. (2015). Gene essentiality and synthetic lethality in haploid human cells. Science 350, 1092–1096.

Staring, J., Von Castelmur, E., Blomen, V.A., Van Den Hengel, L.G., Brockmann, M., Baggen, J., Thibaut, H.J., Nieuwenhuis, J., Janssen, H., van Kuppeveld, F.J.M., et al. (2017). PLA2G16 represents a switch between entry and clearance of Picornaviridae. Nature 541, 412–416.

Staring, J., van den Hengel, L.G., Raaben, M., Blomen, V.A., Carette, J.E., and Brummelkamp, T.R. (2018). KREMEN1 Is a Host Entry Receptor for a Major Group of Enteroviruses. Cell Host Microbe 23, 636–643.e5.

Staring, J., Raaben, M., and Brummelkamp, T.R. (2018). Viral escape from endosomes and host detection at a glance. J. Cell Sci. 131, jcs216259.

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A cknowledgements

En dan het dankwoord. Iets wat mij, net als het afscheid van deze periode en iedereen in het lab, lastiger valt dan ik had verwacht. Na vanaf bijna het begin deel uit te maken van een groep zoals deze, wordt het een hele uitdaging om iets nieuws met dezelfde uitdagingen, creativiteit en opwinding te vinden. Ik heb veel geleden, veel geleerd, veel vriendschappen gesloten en mij altijd uiterst bevoorrecht gevoeld dat ik aan deze reis met de mensen die ik hierna benoem heb mogen maken. Aangezien ik meerdere jaren heb besteed aan dit proefschrift, is het ook niet meer dan logisch dat ik aan een flink aantal mensen dank ben verschuldigd.

Ten eerste Thijn Brummelkamp, mijn begeleider van mijn stage, scriptie en uiteindelijk mijn promotie. Beste Thijn, we hebben er nu bijna 10 jaar opzitten en ik denk dat het eens tijd wordt dat ik op zoek ga naar een andere baas. Ook al hebben we door de jaren heen heel wat gestrubbeld en gestreden, ik heb altijd bijzonder veel respect voor je gehad en heb altijd op je vertrouwd. Ik ken weinig mensen die zo veel passie hebben voor wat ze doen. Je hebt met je creativiteit, intellect en humor een vruchtbare wetenschappelijke omgeving gecreëerd die naar mijn idee uniek is in de

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wereld. Een omgeving die blijkbaar zo fijn is, dat de meeste van je promovendi en postdocs, hun werk met je hebben voorgezet (al is het in je nieuwe bedrijf). Ik wil je bedanken voor het feit dat je mij de mogelijkheid gegeven hebt om een onderdeel te mogen zijn van je lab en ik wens je het allerbeste toe in de toekomst.

Next, I would like to acknowledge my reading committee Emmanuel Wiertz, Jan Carette, Jannie Borst, Anastassis Perrakis, and Frank van Kuppeveld for taking the time to read the manuscript. I would also like to thank my PhD committee:René Bernards, Sjaak Neefjes, and Titia Sixma at the Netherlands Cancer Institute for their supervision throughout the years.

Now, I would like to make an attempt to express my gratitude to everyone I worked with during my PhD. To start with, of course, everyone in the Brummelkamp lab. Guys, I am really, really going to miss you. We all shared the burden of working in a lab with great expectations. We all wanted our BIG paper and we all had to work hard for it. I would like to thank all of you for the group we created together. We (almost) never fought and cracked a lot of jokes (which can never be repeated outside of the lab), we shared everything with everyone (sometimes maybe a bit too much sharing) and were supportive in each other’s endeavors. I am happy that I can consider most of you as my close friends. I never felt any competition within the lab, and I really value the way we supported each other in our (very diverse) research topics. Let me say this again; I am really going to miss all of you.

Vincent Arthur Blomen. Al na twee weken na de start van mijn stage startte ook jij bij Thijn als masterstudent. Ik denk dat ik dan ook wel kan zeggen dat we dit traject samen hebben doorlopen en dat ik het waarschijnlijk niet heb kunnen afronden zonder jouw hulp. Het is voor mij dan ook altijd meer dan vanzelfsprekend geweest dat jij mijn paranimf zou zijn. Aangezien ik in deze sectie van het boek mijn dank behoor uit te spreken, bij deze, bedankt voor je gezelschap, voor je humor en je eindeloze geduld (zelfs wanneer ik voor de grap je computer uitzet zonder iets op te slaan). Ik bewonder je zeer om het feit dat je altijd het geduld op kon brengen om tijd voor iedereen te maken, zelfs wanneer je het echt heel, heel druk had. Bedankt ook, dat ik altijd bij je terecht kon wanneer ik weer eens in de problemen zat. Bedankt voor je hulp bij mijn proeven en het analyseren van mijn screens. Maar bovenal, bedankt voor je vriendschap! Lisa van den Hengel, die enige die haar naam zo goed draagt, dat je nooit een bijnaam hebt gekregen. Ook jij bent iemand die essentieel is geweest in het tot stand komen van mijn promotie. Vanaf het moment

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dat je bij ons bent komen werken, ben je onmisbaar geweest. Je hebt mij enorm (en ik kan dit niet genoeg benadrukken), enorm geholpen met mijn projecten en het was een verademing toen je wat orde in het lab kwam scheppen. Je was altijd bereid om naar mij te luisteren, of het nu om problemen met werk of thuis waren (je had er geld voor moeten vragen!). Bedankt voor het altijd willen doorlezen van mijn mails, manuscripten en zelfs WhatsApp berichten voor die onmisbare spellingscontrole. Ik bewonder je enorm voor je rust en onverstoorbaarheid in je werk en karakter. Ik ga onze koffiepauzes missen, maar ik weet zeker dat we elkaar in de toekomst nog blijven zien. Joppe Daniel Maria Nieuwenhuis, Jopje, we zijn niet tegelijk begonnen, maar wel tegelijk klaar! Ik wens je heel veel succes met je verdediging en met je nieuwe avontuur in Zwitserland. Twee dingen die ik aan je kwijt wil: Denk nog eens aan je “friend-colleague” wanneer je niet meer weet wat je met al je geld moet doen dat je gaat verdienen; en denk maar niet dat je onder je etentjes uit komt met mij en Vincent door naar het buitenland te verhuizen! Matthijs, Mattie, de andere viroloog in het lab. Hoofdstuk 6 was er niet zonder jouw hulp geweest. Daarvoor zeker en zeer mijn dank, maar ik wil je toch vooral bedanken voor je humor en eindeloze goede humeur. Je hebt menigmaal mijn chagrijnige, bijna suïcidale, ochtendhumeur weten te doorbreken met je positiviteit en humor. Markus, Angela Merkel, I started to miss you as soon as you left the lab. You were the best desk buddy someone could ask for and I am happy that you in the end found your way back to the Brummelkamp nest. Lucas, LT, you are the other member of the “Old lab”, and although you left a while ago to pursue your own career in science, you are still missed in the lab. I hope we will see each other again in the future. Elmer, Elmo, ook jij was voor een tijd een bijzondere toevoeging aan ons lab, eerst als masterstudent en later als beginnend promovendus. Ook jou wil ik bedanken voor het werk wat je onze analyse pijplijn hebt gestoken. Ik weet zeker dat je coding kindje Phenosaurus een zonnige toekomst tegemoet gaat. Lisa (2), Astrid, Abdel, and Gian-Luca, the new members of our lab, although our time together was brief, I did appreciate your company and enthusiasm for science. I am quite sure that the lab is in safe hands with you guys. Also, special thanks for the other lab members Sjoerd and Ferdy, Bianca, Gregor, Jan Pruszak, Chong, and the students that worked in the lab over the years and of which some have become true friends: Nicolaas, Jeff, Peter, Ana, Shuai, en natuurlijk mijn eigen student Max. Ook al moest ik je van Matthijs stelen om aan een student te komen, het was meer dan de moeite waard. Onze gesprekken waren een alleraardigste afleiding van de dagelijkse sleur in het lab en ik hoop dat ook jij met succes je promotietraject kunt doorlopen in het AMC.

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Many thanks to my collaborators Tassos, and Elli. Tassos, thank you for your patience and willingness to postpone the publication of our manuscript again and again, thereby giving me the time to solve the biggest puzzle of my life. Frank, ook jou wil ik bedanken voor de samenwerking op verschillende picornavirus projecten. Samen met Jim en Hendrik Jan hebben jullie mij een kijkje kunnen geven in een echt virus lab. Jullie enthousiasme voor de projecten was eindeloos en ik heb ontzettend veel van jullie geleerd.Jan , ik heb je al bedankt voor de moeite die je hebt genomen om plaats te nemen in mijn promotiecommissie, maar ik wil je ook graag nog bedanken voor onze tijd in Boston. Als mijn dagelijkse stagebegeleider heb ik daar ontzettend veel van je geleerd en ik ben je dankbaar voor je interesse die je door de jaren heen altijd hebt getoond voor mijn projecten. Malini Varadarajan, your Dutch must be rusty (by now), so let me say this in English. Although we haven’t spoken with each other for a few years, I am still grateful for your guidance and friendship when I just arrived in a new and foreign country. I will never forget your kindness.

Thank you, to everybody else that I had the pleasure of working with on B8:Tati, for always helping out when we needed you, Hans, voor je prachtige EM figuren die zich een weg hebben weten te vinden in hoofdstuk 3, Patrick, and Magda, also your help was indispensable for the publications in this thesis. Danny, Robbert, Willem Jan and Michael, you all left already, but I always enjoyed our gossip sessions. Many thanks to the other members of B8: Luca, Andrea, Nassos, Thanga, Shreya, Yvette, Doreth, Yoshi, Fernando, Robbie, Wouter, Roy, Susanne, Bart, Misbha, Xiaohu, Foteini en uiteraard het hoofd van onze afdeling Titia. In het bijzonder wil ik graag de mensen van B8 (B7 en C2) bedanken die al die jaren voor mij gezorgd hebben. Mirna, Caroline, Patty, Pim en Herrie zonder jullie luisterend oor en administratieve ondersteuning was er nooit iemand gepromoveerd op B8. Hetzelfde geldt voor Mohammed, jij hebt zonder klagen en altijd vrolijk er persoonlijk voor gezorgd dat ons lab niet in totale chaos is vervallen. Dank aan alle mensen die ik door de jaren heen heb leren kennen van de facilitaire afdelingen binnen het NKI. Lenny, Lauran en Marjolijn, ik heb zoveel tijd doorgebracht in die donkere kamers achter de confocal dat ik er waarschijnlijk nog jaren van blijf dromen, maar jullie hulp en vriendelijkheid heeft het zeker een stuk dragelijker gemaakt. Ook Desiree is tot het einde bijzonder behulpzaam geweest. Jij dan ook bedankt voor je onvermoeibare inzet voor de wetenschap en de wetenschappers binnen het NKI. Al ben ik al jaren niet meer in het muizenhuis geweest, ik weet dat er een groot team van dierverzorgers dag en nacht aan het werk is om voor onze dieren te zorgen. Al

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het contact dat ik met jullie gehad heb is altijd vriendelijk en prettig geweest. Mijn dank aan jullie is dan ook groot.

To all the others that kept my life bearable during my time at the NKI. Tessy, Daniela, Chiara, and Jonne. Thank you for your company and friendship during the years. Discussing my science/life problems with you kept me going until the very end. Hetzelfde gaat op voor David en Julia. Ik ben jullie zeer dankbaar voor de eindeloze discussies over experimenten, manuscripten, figuren, modellen en vervelende editors, maar zeker ook voor het feit dat ik altijd naar B6 kon “vluchten” wanneer ik het even niet meer zag zitten. Ik ben dankbaar voor jullie nuchtere kijk op het leven en jullie vriendschap en kijk met vertrouwen naar jullie voortgang in jullie promotie trajecten. Thanks to everyone else that helped out in some way or another: Jeroen, Lorenzo, Justina, Rui, Michiel, Femke, Judith, Tao, Aniruth, Ricardo, Bastiaan, Boekie, Maarten, Georgi from Georgia, Sanne, Hilje, Henri en René Medema.

Dan toch nog even terug naar mijn opleiding. Geert Jan van Tetering, mijn allereerste stagebegeleider. Van jou heb ik de passie voor de wetenschap geleerd en ik ben je daarvoor dankbaar. Marc Vooijs, op jouw advies en met jouw hulp ben ik naar Thijn in Boston vertrokken. Ik weet niet of dit het plan was wat je in gedachten had, maar het heeft mijn leven compleet veranderd.O ok hier ben ik dankbaar voor.

Mijn vrienden en familie. Ik heb mij door velen van jullie door de jaren heen gesteund gevoeld, maar ik wil een paar van jullie in het bijzonder bedanken. Judith, Snuud, je bent mijn liefste huisgenootje en samen met je zus Helene, mijn Nel, hebben jullie mij onophoudelijk gesteund en mij door mijn donkerste tijden heen geholpen. Hetzelfde geldt voor de rest van mijn tweede familie, de familie Karels. Dank dus ook aan Joost, Marcel B, Johan, en Marleen. Ook Lisette en Bas, bedankt dat ik jaren aan jullie keukentafel heb mogen vertellen over mijn wetenschappelijke avonturen. Ook dank aan mijn andere meiden; Annemarie, Jorien en Sonja en natuurlijk de rest van mijn Alphense clan.

Ik ben oneindig dankbaar voor mijn eigen familie. Judith, mijn lieve nicht, je hebt altijd interesse getoond in mijn werk en ook al spreken we elkaar niet vaak, ik waardeer je steun enorm. Mijn lieve schoonzusje en moeder van mijn neefje Daan, Annemiek, jouw liefde, steun en trouw aan je familie heb ik altijd gevoeld en zijn oprecht inspirerend.

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Vader Gert en Moeder Ans Staring, bij deze de mogelijkheid voor mij om mijn dankbaarheid naar jullie uit te spreken. Ook al hebben jullie mijn keuzes niet altijd begrepen en kwam de wereld van de wetenschap jullie soms wat vreemd over, toch hebben jullie mij altijd gesteund en ondersteund toen ik dat het meest nodig had. Ik prijs mij gelukkig met ouders die zo een stabiel en veilig thuis hebben gecreëerd waarvan ik weet dat ik er altijd naar terug kan keren.

Dan rest mij alleen nog een dankbetuiging aan mijn andere paranimf, mijn liefste broer, Marcel. Je bent altijd mijn grote broer geweest waar ik bij terecht kon. Ook van jou heb ik bijzonder veel steun genoten. Jij bent een van de weinige mensen waar ik mij nooit veroordeeld voel in de keuzes die ik maak. Door mij aan te moedigen in mijn keuzes, ook wanneer ik twijfelde, heb jij er menig maal voor gezorgd dat ik een weg in ben geslagen die andere misschien wat vreemd of risicovol vonden. Hierdoor ben ik de persoon geworden die ik ben. Ik zal je daar altijd dankbaar voor zijn. Je bent voor mij een klankbord die mij enorm waardevol is. Het is mij dan ook een eer dat je naast mij wilt staan tijdens de verdediging van dit proefschrift.

Het was een hele reis en dit is waar het eindigt

Jacqueline

PS_JS_def.indd 174 08-10-18 08:47 Roll the dice

if you’re going to try, go all the way. otherwise, don’t even start. if you’re going to try, go all the way. this could mean losing girlfriends, wives, relatives, jobs and maybe your mind. go all the way. it could mean not eating for 3 or 4 days. it could mean freezing on a park bench. it could mean jail, it could mean derision, mockery, isolation. isolation is the gift, all the others are a test of your endurance, of how much you really want to do it. and you’ll do it despite rejection and the worst odds and it will be better than anything else you can imagine. if you’re going to try, go all the way. there is no other feeling like that. you will be alone with the gods and the nights will flame with fire. do it, do it, do it. do it. all the way all the way. you will ride life straight to perfect laughter, its the only good fight there is.

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