Influenza a Virus

Dan Dou Influenza A Virus

Influenza A Virus Spatial analysis of influenza genome trafficking and the evolution of the neuraminidase protein

Dan Dou

To: NA

ISBN 978-91-7797-885-5

To: NA NA To: NA To:

NA Department of Biochemistry and Biophysics NA To: NA To: To:

Doctoral Thesis in Biochemistry at Stockholm University, Sweden 2019 Influenza A Virus Spatial analysis of influenza genome trafficking and the evolution of the neuraminidase protein Dan Dou Academic dissertation for the Degree of Doctor of Philosophy in Biochemistry at Stockholm University to be publicly defended on Monday 2 December 2019 at 10.00 in Vivi Täckholmsalen (Q-salen), NPQ-huset, Svante Arrhenius väg 20.

Abstract Influenza A viruses (IAVs) are a common infectious agent that seasonally circulates within the human population that causes mild to severe acute respiratory infections. The severity of the infection is often related to how the virus has evolved with respect to the pre-existing immunity in the population. For IAVs, the most common mechanisms to avoid the immune response are to vary the surface antigens, hemagglutinin (HA) and neuraminidase (NA), by processes known as antigenic drift and shift. Antigenic drift refers to point mutations that accumulate in HA and NA as a result of the antibody-mediated selection pressure that exists in the population. The majority of the changes attributed to antigenic drift localize to HA and NA surface exposed regions, however this does not exclude that drift can also result in the selection of residues that are not exposed. One region where non-exposed residues have potentially been selected for is the NA transmembrane domain (TMD) of human H1N1 IAVs, where a temporal bias exists for the accumulation of polar residues. By examining these sequence changes in the NA TMD, we found that the polar residues contribute to the amphipathic characteristic of the NA TMD, which mediates the oligomerization of the N-terminus. As more polar residues became incorporated, the strength of the TMD- TMD interaction increased, presumably to benefit the NA head domain assembly into a functional tetramer. We determined that the amphiphilic drift in the NA TMD is able to bypass the strict hydrophobicity required for membrane insertion at the endoplasmic reticulum because it can utilize the co-translational translocation process to facilitate the insertion and inversion of its non-ideal TMD. The contribution of the TMD to proper NA assembly was traced to the formation of the Ca2+ binding pocket that is located at the center of the tetrameric assembly, as this pocket lies above the stalk linker regions and must be occupied for NA to function. In addition to antigenic drift, NA and HA can also undergo antigenic shift. Antigenic shift occurs when either of the gene segments encoding NA or HA are exchanged with ones from another IAV encoding another subtype of NA or HA. Different from antigenic drift, antigenic shift can only occur when a cell is co-infected and most investigations on the process of reassortment have been made at the protein level due to the methodological issues for labeling the RNA genome in situ. To overcome these technical limitations, we developed an in situ RNA labeling approach that provides highly specific spatial resolution of the IAV genome throughout the infection process. By applying this approach to temporally analyze the co- infection process, we found that the entry of a second IAV is stalled in the cytoplasm if another IAV has begun to replicate. Together, these results provide insight into the low frequency of antigenic shift in nature and provide evidence that non- exposed residues may make an underappreciated contribution to NA antigenic drift in human H1N1 viruses.

Keywords: Influenza A virus, IAV, neuraminidase, NA, IAV genome trafficking, viral entry, viral replication, co- infection, antigenic drift, antigenic shift, NA assembly, transmembrane domain, evolution.

Stockholm 2019 http://urn.kb.se/resolve?urn=urn:nbn:se:su:diva-175202

ISBN 978-91-7797-885-5 ISBN 978-91-7797-886-2

Department of Biochemistry and Biophysics

Stockholm University, 106 91 Stockholm

INFLUENZA A VIRUS

Dan Dou

Influenza A Virus

Spatial analysis of influenza genome trafficking and the evolution of the neuraminidase protein

Dan Dou ©Dan Dou, Stockholm University 2019

ISBN print 978-91-7797-885-5 ISBN PDF 978-91-7797-886-2

Printed in Sweden by Universitetsservice US-AB, Stockholm 2019 To my dear mother,

I would have never gotten this far without you.

List of publications

I. Polar Residues and Their Positional Context Dictate the Transmem- brane Domain Interactions of Influenza A Neuraminidases. Nordholm J, da Silva DV, Damjanovic J, Dou D, Daniels R. J Biol Chem. 2013 Apr 12;288(15):10652-60. doi: 10.1074/jbc.M112.440230

II. Type II Transmembrane Domain Hydrophobicity Dictates the Co- translational Dependence for Inversion. Dou D, da Silva DV, Nordholm J, Wang H, Daniels R. Mol Biol Cell. 2014 Nov 1;25(21):3363-74. doi: 10.1091/mbc.E14-04-0874

III. Structural Restrictions for Influenza Neuraminidase Activity Promote Adaptation and Diversification. Wang H, Dou D, Östbye H, Revol R, Daniels R. Nat Microbiol. 2019 Aug 26. doi: 10.1038/s41564-019-0537-z

IV. Analysis of IAV Replication and Co-infection Dynamics by a Versa- tile RNA Viral Genome Labeling Method. Dou D, Hernández-Neuta I, Wang H, Östbye H, Qian X, Thiele S, Resa-Infante P, Kouassi NM, Sender V, Hentrich K, Mellroth P, Hen- riques-Normark B, Gabriel G, Nilsson M, Daniels R. Cell Rep. 2017 Jul 5;20(1):251-263. doi: 10.1016/j.celrep.2017.06.021.

Publications not included in this thesis

I. The Influenza Virus Neuraminidase Protein Transmembrane and Head Domains Have Coevolved. da Silva DV, Nordholm J, Dou D, Wang H, Rossman JS, Daniels R. J Virol. 2015 Jan 15;89(2):1094-104. doi: 10.1128/JVI.02005-14

II. Translational Regulation of Viral Secretory Proteins by The 5' Cod- ing Regions and a Viral RNA-binding Protein. Nordholm J, Petitou J, Östbye H, da Silva DV, Dou D, Wang H, Dan- iels R. J Cell Biol. 2017 Aug 7;216(8):2283-2293. doi: 10.1083/jcb.201702102

III. Influenza A Virus Cell Entry, Replication, Virion Assembly and Movement. Dou D, Revol R, Östbye H, Wang H, Daniels R. Front Immunol. 2018 Jul 20;9:1581. doi: 10.3389/fimmu.2018.01581

Abstract

Influenza A viruses (IAVs) are a common infectious agent that seasonally circulates within the human population that causes mild to severe acute respiratory infections. The severity of the infection is often related to how the virus has evolved with respect to the pre-existing immunity in the population. For IAVs, the most common mechanisms to avoid the immune response are to vary the surface antigens, hemagglutinin (HA) and neuraminidase (NA), by processes known as antigenic drift and shift.

Antigenic drift refers to point mutations that accumulate in HA and NA as a result of the antibody-mediated selection pressure that exists in the population. The majority of the changes attributed to antigenic drift localize to HA and NA surface exposed regions, however this does not exclude that drift can also result in the selection of residues that are not exposed. One region where non-exposed residues have potentially been selected for is the NA transmembrane domain (TMD) of human H1N1 IAVs, where a temporal bias exists for the accumulation of polar residues. By examining these sequence changes in the NA TMD, we found that the polar residues contribute to the amphipathic characteristic of the NA TMD, which mediates the oligomerization of the N- terminus. As more polar residues became incorporated, the strength of the TMD-TMD interaction increased, presumably to benefit the NA head domain assembly into a functional tetramer. We determined that the amphiphilic drift in the NA TMD is able to bypass the strict hydrophobicity required for membrane insertion at the endoplasmic reticulum because it can utilize the co- translational translocation process to facilitate the insertion and inversion of its non-ideal TMD. The contribution of the TMD to proper NA assembly was traced to the formation of the Ca2+ binding pocket that is located at the center of the tetrameric assembly, as this pocket lies above the stalk linker regions and must be occupied for NA to function.

In addition to antigenic drift, NA and HA can also undergo antigenic shift. Antigenic shift occurs when either of the gene segments encoding NA or HA are exchanged with ones from another IAV encoding another subtype of NA

or HA. Different from antigenic drift, antigenic shift can only occur when a cell is co-infected and most investigations on the process of reassortment have been made at the protein level due to the methodological issues for labeling the RNA genome in situ. To overcome these technical limitations, we developed an in situ RNA labeling approach that provides highly specific spatial resolution of the IAV genome throughout the infection process. By applying this approach to temporally analyze the co-infection process, we found that the entry of a second IAV is stalled in the cytoplasm if another IAV has begun to replicate. Together, these results provide insight into the low frequency of antigenic shift in nature and provide evidence that non-exposed residues may make an underappreciated contribution to NA antigenic drift in human H1N1 viruses.

Contents

Overview of Influenza A virus ...... 1 The composition of IAV ...... 1 IAV subtypes ...... 2 Influenza life cycle ...... 4 Binding and Fusion of IAVs ...... 4 Nuclear import of vRNPs...... 5 Viral mRNA transcription ...... 7 Replication and assembly of vRNPs ...... 8 vRNP export and plasma membrane targeting ...... 9 Membrane protein synthesis and maturation ...... 11 Protein targeting to the ER ...... 11 Transmembrane insertion into the ER membrane ...... 12 Protein maturation within the ER lumen ...... 14 Structure and function of NA ...... 17 Structure of NA ...... 17 Function of NA ...... 18 Antibodies against NA ...... 21 Results summary ...... 24 Conclusions and future perspective ...... 27 Sammanfattning på svenska ...... 30 Acknowledgments ...... 30 References ...... 32

Abbreviations

ADCC antibody-dependent cell-mediated cytotoxicity ADP antibody-dependent phagocytosis CAS cellular apoptosis susceptibility protein ER endoplasmic reticulum HA hemagglutinin IAV Influenza A virus LMB leptomycin B M1 matrix protein 1 M2 matrix protein 2 NA neuraminidase NES nuclear export sequence NK natural killer NLS nuclear localization sequence NPC nuclear pore complex NP nucleoprotein PA polymerase acidic protein PB1 polymerase basic protein 1 PB2 polymerase basic protein 2 SRP signal recognition particle SS ER signal sequence TMD transmembrane domain vRNP viral ribonucleoprotein vRNA viral RNA

Overview of Influenza A virus

Influenza A viruses (IAVs) are one of the most common contagious path- ogens that circulate in the human population yearly. Globally, there are two forms of influenza that circulate: seasonal epidemic influenza and sporadic pandemic influenza [1]. Depending on the viral strain, pre-existing immunity and the immune response of the host, influenza viruses can cause mild asymp- tomatic illnesses to severe acute respiratory infections. In humans, IAVs mainly infect the respiratory epithelial cells, but to do so the virus first needs to penetrate the mucosal barrier to reach the surface of the epithelium where it can initiate the infection process by attaching to cell receptors that trigger its internalization. Once inside the cell, the replication and assembly of progeny IAVs is a complicated process that requires coordination between the replication of the viral genome and the synthesis of the viral proteins. This coordination is especially complex for IAVs because the replication of the genome in the host cell nucleus is coupled to the synthesis of the viral proteins in the cytosol and at the endoplasmic reticulum [2]. To date, many of the mechanisms that assist in the coordination of the IAV replication process have been identified. However, the properties that define the negative selection pressure on NA and HA remain to be established due to the limited knowledge of the maturation and trafficking requirements for the different subtypes of these important surface antigens.

The composition of IAV

IAVs have a complex organization that includes a lipid envelope surround- ing eight RNA gene segments, which encode for one or more viral proteins [3]. [Fig. 1A] The viral envelope is a host-derived lipid bilayer in which the viral membrane proteins hemagglutinin (HA), neuraminidase (NA) and matrix protein 2 (M2) are embedded [4, 5]. HA and NA are glycoproteins with HA being more abundant. The ratio between HA and NA on the viral surface can vary from 5:1 to 2:1 [6], and M2 is the least abundant of the three. Underneath

the viral envelope, the matrix protein 1 (M1) forms a protein layer that connects the viral gene segments to the envelope. Each of the negative-sense single-stranded viral RNA (vRNA) gene segments are packed into separate viral ribonucleoprotein (vRNP) complexes where the vRNA is bound by multiple copies of the viral nucleoprotein (NP) and a single copy of the viral polymerase that consists of PB1, PB2 and PA [7, 8]. NP associates with 12 nucleotide stretches in the vRNA with a partial G bias, whereas the polymerase binds to the helical hairpin “panhandle” structure that is formed by the partial anneal- ing of the complementary 5’ and 3’ ends of the vRNA [9, 10] [Fig. 1B].

Gene A segments B IAV PB2 M2 PB1 vRNP NA PA PB1 HA HA PA 3’ NP PB2 5’ NA vRNA NP M1 M M1 M2

spliced genes NS NS1 vRNPs NS2 Figure 1. Schematic of an influenza A virus (IAV). (A) Diagram of an IAV particle and its viral RNA (vRNA) gene segments. The viral membrane proteins HA, NA and M2 are shown on the viral surface. The M1 protein is depicted on the luminal side of the virion together with the eight viral ribonucleoprotein complexes (vRNPs). (B) Di- agram of a vRNP. A single-stranded viral RNA (vRNA) is wrapped around multiple copies of NP. The terminal 5’ and 3’ regions are partially base paired and bound by the heterotrimeric vRNA-dependent RNA polymerase (PB2, PB1, PA) via PB1.

IAV subtypes

IAVs are classified by the genetic and antigenic properties of the two surface antigens HA and NA into subtypes. Sixteen influenza HA subtypes (H1 to H16) and nine NA subtypes (N1 to N9) have been identified in aquatic birds, the natural reservoir of IAV [11]. Of these, only three IAV subtypes have caused pandemics and subsequent seasonal epidemics in the human population, which are H1N1 (1918 Spanish influenza and 2009 swine influenza), H2N2 (1957 Asian influenza), and H3N2 (1968 Hong Kong influenza) [12]. Similar to many other viruses, the IAV surface antigens evolve constantly to escape the host immune response. This evolution is assisted by the error prone

viral RNA polymerase, which lacks a proof-reading function resulting in error rates that have been estimated to be about 10^-5 to as high as 10^-4 [13-17]. These errors can lead to amino acid changes and alterations to the protein structure. When the virus is subject to pressure from the immune system, substitutions in HA and NA that promote immune system escape are often selected for over time, which is known as antigenic drift [18].

Due to the segmentation of the genome, IAVs can also exchange segments between different viruses through a process called reassortment. In contrast to substitutions, reassortment requires that a cell is co-infected by the different IAVs so the gene segments can mix during the assembly process and produce progeny with different arrays of the parental IAV gene segments. When the exchange occurs between either HA or NA, a new IAV subtype can appear. This process is known as antigenic shift. In the past, three major human IAV pandemics have occurred due to antigenic shift. The 1957 H2N2 and 1968 H3N2 pandemics both were caused by a novel HA entering into the human population by reassortment, whereas the 2009 H1N1 pandemic of swine origin resulted from the incorporation of a novel subtype 1 NA [12, 19, 20].

Influenza life cycle

Viruses rely on host cells for replication and the synthesis of progeny viruses. The main challenge for viruses is to deliver the genetic material to the correct place in the cell to gain access to the necessary cellular machinery for their replication. As IAV is an enveloped virus that replicates in the nucleus, IAV proteins and gene segments need to cross several membrane barriers during the viral life cycle. To accomplish this feat, IAVs depend heavily on the host machinery and pathways due to the small genome size and the limited number of viral gene products it encodes.

Binding and Fusion of IAVs

The initial steps of an IAV infection involves binding, endocytosis and fusion, which are primarily mediated by surface glycoprotein HA. During replication HA is expressed as a pro-protein that is proteolytically processed into the subunits HA1 and HA2 [21]. HA1 contains a receptor binding pocket that can bind to host cell surface glycoconjugates via terminal sialic acid residues [5, 22], whereas HA2 contains the fusion peptide that exposes upon pH changes. Upon reaching the cell surface, the HA1 receptor binding domain associates with the surface receptors. The binding triggers endocytosis of the virus either through a clathrin-dependent [23, 24], or a macropinocytosis pathway [25] [Fig. 2(i)], both of which enable the virus to enter the host cell. Once inside the cell, the pH of the virus containing vesicle decreases through the vacuolar-type proton ATPase, facilitating the vesicle to mature to a late endosome [26]. As a consequence of the low pH inside the endosome, the viral M2 ion channel opens, allowing an influx of protons that acidifies the inside of the virion and causes the dissociation of M1 from the vRNPs [27, 28]. The low pH also triggers a conformational change of HA, which exposes the fusion peptide in the HA2 subunit [Fig. 2(ii)]. Upon exposure, the N-terminal domain of the fusion peptide can insert into the endosomal membrane, while the C- terminal domain of HA2 is anchored in the viral envelope [Fig. 2(iii)] [29].

Once in this pre-hairpin conformation, the HA2 trimer then folds back on itself positioning the endosomal membrane close to the viral envelope. When the two membranes become close enough, the lipids reorganize, begin to mix, and ultimately form a fusion pore [Fig. 2(iv)] [30], which enables the release of the vRNPs into the cytoplasm [Fig. 2(v)].

Endosome M2 Cell HA pH 5 (i) pH 5 (ii) (iii) (iv) IAV pH 5

(v)

HA HA HA ‘receptor’ binding Figure 2. Cell entry of IAVs. (i) IAVs initiate cell entry by attaching to the cell surface via HA binding to a sialylated receptor that facilitates the endocytosis of the virus. (ii) The endocytic vesicle matures to an endosome via a decrease in pH. This low pH triggers a conformational change in HA that exposes the fusion peptide for insertion into the endosomal membrane and it also causes the M2 ion channel to open. (iii) Opening of the M2 channel acidifies the interior of the virus causing the vRNPs to release from M1. (iv) Following the formation of the pre-hairpin conformation), the HA helical bundle collapses into a trimer of hairpins resulting in the formation of the fusion pore between the viral and endosomal membranes. (v) vRNPs are released through the fusion pore into the cytosol. The illustration is modified from Dou et al., 2018.

Nuclear import of vRNPs

The nucleus is a dual-membrane enclosed organelle that uses the nuclear pore complex (NPC) to regulate the transport of molecules across the membrane. Small molecules can passively diffuse through the NPC, whereas molecules above ~40 kDa have to be transported via an energy-dependent pathway [31, 32]. To replicate the viral genome, IAVs need to deliver each vRNP to the nucleus. Since the vRNPs are large complexes composed of the vRNA, a single copy of the viral polymerase and numerous copies of NP, IAVs are reliant on the energy-dependent nuclear transport pathway require to facilitate the import of the vRNPs into the nucleus [33, 34].

The classical nuclear import machinery contains two adaptor proteins including importin-a and importin-b, RanGTPase and the cellular apoptosis susceptibility protein (CAS). This machinery recognizes cargo that contains a nuclear localization sequence (NLS) and is responsible for directing it to the nucleus. A typical NLS is a sequence rich in arginine and lysine residues, which can be located anywhere in the amino acid sequence of a protein [35]. The NP protein, the major component of a vRNP, has an exposed N-terminal NLS that enables importin-a to recognize a vRNP as a cargo protein [36-38]. Binding of importin-a to the NLS initiates the nuclear import process as it results in the recruitment of importin-b. Importin-b associates with importin-a, forming the importin-a/importin-b-cargo complex [Fig. 3(i)] that is necessary for translocation of the cargo into the nucleus [Fig. 3(ii)]. Once the complex reaches the nucleus, the cargo protein is released sequentially. Importin-b dissociates first by interacting with RanGTPase [Fig. 3(iii)] after which importin- a dissociates via the cellular CAS protein [Fig. 3(iv)]. Importin-a and importin-b are then individually recycled back to the cytoplasm, while the cargo remains in the nucleus [Fig. 3(v)] [32]. Incoming vRNP Cytoplasm Nucleus PB1 NLS PA 3’

PB2 5’ NPC vRNA NP (v) (i) (v) (iv) RanGDP (iii)

(ii)

Importin a RanGTP Importin b CAS RanGTP Figure 3. Nuclear import of a vRNP. (i) Importin-a recognizes the nuclear localization sequence (NLS) on NP and recruits importin-b. (ii) The vRNP-importin-a-importin-b complex is then transported through the nuclear pore complex (NPC). (iii) Importin-b dissociates from the complex by interacting with RanGTP. (iv) The vRNP is released from importin-a which associates with the nuclear export factor CAS RanGTP. (v) Importin-a and importin-b are exported to the cytoplasm and the hydrolysis by RanGTP to RanGDP facilitates the release of importin a and importin b for another import cycle.

Viral mRNA transcription

Eukaryotic mRNA is transcribed from DNA by a RNA polymerase and is then subject to processing by several different enzymes in the nucleus to produce a mature mRNA. The mature mRNAs generally possess a 5' 7-methyl- guanosine cap (5’ m7G cap) and a 3’ polyadenyl moiety. As IAVs are dependent on the host translational machinery for viral protein synthesis, the viral mRNAs, although transcribed by viral RNA-dependent RNA polymerase, need to mimic the structure of a cellular mRNA.

The viral mRNA transcription process begins once the vRNPs arrive in the nucleus [Fig 4. (i)]. The viral mRNA obtains its 5’ m7G cap from host mRNAs through a mechanism called ‘cap snatching’ [39]. This process is initiated by the cap-binding domain of the PB2 subunit [40]. Association with the 5’ host mRNA cap allows the endonuclease domain of the PA subunit to cleave off the cap 10-13 nucleotides downstream [41]. This capped mRNA stretch is re- positioned to the PB1 catalytic center, where the 3’ end base-pairs with the 3’ end of vRNA. Then PB1 uses the capped mRNA as a primer for synthesizing viral mRNA using vRNA as a template [42]. After transcription has been initiated, the 5’-cap dissociates from the PB2 cap-binding domain and likely binds the host nuclear cap-binding complex, allowing for nuclear export of the final product [43, 44]. The 3’ polyadenylated tail is acquired by a reiterative stuttering process, where the polymerase repeatedly transcribes the short polyU sequence located at the vRNA 5’ end [45].

As for cellular mRNAs, viral mRNAs are exported to the cytoplasm through the NPC [Fig. 4(ii)]. Once in the cytoplasm, the ribosome recognizes newly synthesized mRNA and protein synthesis is initiated. All of the soluble viral proteins (PB2, PB1, PA, NP, NS1, NS2, and M1) are translated by cytosolic ribosomes, whereas the membrane proteins (HA, NA and M2) are synthesized on the ER membrane. The newly synthesized viral proteins (PB2, PB1, PA and NP) are imported to the nucleus for vRNP replication and assembly [Fig. 4(iii)]. PB2, PA, and NP each contain a NLS that can use the cellular importin-a/importin-b pathway for nuclear import, whereas PB1 is transported through its association with PA in the cytoplasm [37, 46]. HA, NA and M2 synthesis will be discussed in the ‘Membrane protein synthesis and maturation’ section.

Replication and assembly of vRNPs

Replication of the viral genome can be separated into two major steps. First, the negative-sense vRNAs are transcribed to a complimentary positive-sense RNA (cRNA). Second, cRNAs serve as a template for additional progeny vRNA synthesis.

Ribosome A(n) NA HA M2

(ii) NP PA PB1 PB2

NPC (iii) A(n) ER splicing A(n) A(n) mRNA(+) mRNA(+) x 8 x 8 (v) (vii) (i) (viii)

(iv) (vi) (x) (ix) Incoming cRNP(+) Progeny vRNP(-) x8 vRNP(-) x8 Nucleus Cell

Figure 4. Viral mRNA transcription, replication and assembly of progeny vRNPs. (i) Viral mRNAs are transcribed in the nucleus from incoming vRNPs by the attached RNA-dependent RNA polymerase. (ii) The mRNAs are exported to the cytoplasm for protein synthesis. (iii) Newly synthesized NP, PA, PB1 and PB2 are targeted to the nuceus by the importin a-importin b pathway. (iv) cRNAs are transcribed from vRNAs and (v) assemble to cRNPs together with newly synthesized viral proteins. (vi) cRNPs can further be used to synthesize progeny vRNA and (vii) assemble to progeny vRNPs with de novo viral proteins. The progeny vRNP can continue to transcribe more viral mRNA (vii), replicate cRNA (ix), or export from the nucleus together with other viral proteins (x). The illustration is modified from Dou et al., 2018.

The transcription of cRNA is a primer-independent process that uses the viral polymerase from the incoming vRNP template[47]. Newly synthesized cRNA associates with newly synthesized NP once it exits from the polymerase exit tunnel [Fig. 4(iv and v)][48, 49]. Each NP monomer binds to G-rich stretches with poor U-bias, about 12 nucleotides long. As for vRNPs, multiple

copies of NP, together with the cRNA, assemble into a cRNP complex [50, 51]. The final cRNP is formed when newly synthesized viral polymerases associate at the 5’-3’ panhandle structure of the cRNA. This resident polymerase is then able to use the cRNA to produce new vRNAs [Fig. 4(vi)]. Following vRNA transcription, newly synthesized viral proteins assemble in a similar way as during cRNP assembly [Fig. 4(vii)] [52]. Newly formed vRNPs can then be used for further transcription [Fig. 4(viii)], replication [Fig. 4(ix) or can be transported from the nucleus to the plasma membrane for incorporation into progeny viruses [Fig. 4(x)]. vRNP export and plasma membrane targeting

The cellular chromosomal maintenance 1 (CRM1, also known as exportin 1)-dependent nuclear export pathway is used to actively transfer proteins with a leucine-rich nuclear export sequence (NES) from the nucleus to the cytoplasm. CRM1 binds to the cargo protein via its NES, and associates with RanGTPase, which transports the cargo across the NPC. At the cytoplasmic side RanGTP is hydrolyzed, which facilitates the cargo protein release. Ran- GDP can then cycle back to the nucleus, undergo nucleotide exchange and be used in the next round of nuclear export [32].

IAVs have been shown to use this pathway, as the inhibition of the CRM1- dependent pathway, by the inhibitor leptomycin B (LMB), causes a major portion of vRNPs to remain in the nucleus [53, 54]. However, as none of the proteins forming the vRNP contain a NES, an addition viral protein is needed to facilitate the export of the genome. This protein is NS2, also known as NEP (nuclear export protein). Soluble NS2, as well as M1, are imported into the nucleus during replication to facilitate vRNP export [Fig. 5(i)]. M1 is thought to function as an adaptor protein that links NS2 to the vRNP. Through the NS2 NES, CRM1 is able to associate with the vRNP-M1-NS2 complex and export it to the cytoplasm [Fig. 5(ii)]. In the cytoplasm, NS2 disassociates from the vRNP, while M1 stays attached, possibly to prevent re-targeting of the vRNP back to the nucleus by blocking the NLSs on the associated NP molecules.

After nuclear export, the vRNPs continue to traffic towards the plasma membrane by interacting with Rab-11 [Fig. 5(iii)]. Rab-11 is a small GTPase involved in trafficking vesicles between the trans-Golgi network, recycling

endosomes, and the plasma membrane [55]. vRNPs associate with Rab-11 via the polymerase subunit PB2 [56]. This could be a mechanism used to make sure that the newly assembled vRNPs carry a polymerase complex. There are two models attempting to describe how vRNPs are trafficked in the cytoplasm. The traditional view is that Rab-11 acts as an adaptor that connects vRNPs to the recycling endosome, and these endosomes then use microtubules to traffic towards the cell surface [Fig. 5(iiia)] [56-58]. A recently proposed model sug- gests that IAV infections cause tubulation of the ER membrane network [Fig.

5(iiib)]. The vRNPs would then associate on the ER via Rab-11 and be transported through this tubular ER network towards the plasma membrane [59]. Once the vRNPs reach the plasma membrane, all eight of the different vRNPs incorporate into the virion together with the other viral membrane proteins. However, how vRNPs are transferred from the vesicles, or the ER network, to the plasma membrane remains unclear.

Ribosome Cell A(n) Modified ER ? ER

NS2 M1 (i) (iiib)

CRM-1 Ran-GTP (ii) (iiia) MTOC

CRM-1 microtubule Nucleus Ran-GDP NS2 Rab 11

Figure 5. vRNP export from the nucleus and trafficking towards the plasma membrane. (i) Newly synthesized M1 decorates vRNPs via interaction with NP. NS2 plays a role as an adaptor protein between M1 and the CRM-1 protein. (ii) The CRM-1- vRNP complex is exported from the nucleus by Ran-GDP/GTP. (iii) The vRNPs then associate with vesicles via Rab-11 and are trafficked towards the plasma membrane through either microtubules (iiia) or a modified ER system (iiib). The illustration is modified from Dou et al., 2018.

Membrane protein synthesis and maturation

Not only vRNPs, but also membrane proteins, need to reach the plasma membrane for the assembly of progeny viral particles. To achieve this localization, the three IAV membrane proteins, HA, NA and M2, utilize the cellular secretory pathway.

Protein targeting to the ER

For ER-targeting, proteins generally possess a signal sequence (SS), which is a stretch of 13 to 36 amino acids located at the N-terminus of the protein. Typically, a SS contains a stretch of 10 to 15 hydrophobic amino acid with one or more positively charged residues at the upstream and a signal peptidase cleavage site at the downstream site [60]. Whereas HA contains a typical SS at its N-terminus [Fig 6A.], NA and M2 use their N-terminal transmembrane domain (TMD) for targeting to the ER [Fig. 6B, C]. During protein translation, the SS is the first part of the protein to emerge from the ribosome exit tunnel [Fig. 6(i)]. The signal recognition particle (SRP), which constantly scans newly synthesized polypeptides for the presence of a SS, recognizes the SS, associates with it in a GTP-bound state and pauses the ribosome translation [Fig. 6(ii)]. The SRP directs the ribosome-nascent chain complex to the GTP- bound SRP receptor that is located next to the Sec61 translocon. GTP-bound SRP and GTP-bound SRP receptor form a heterodimer complex and the two GTPases reciprocally activate the GTP hydrolysis activities in each other. Hy- drolysis of GTP in SRP and in the SRP receptor leads to the transfer of the ribosome-nascent chain complex to the Sec61 translocon and the elongation of the polypeptide chain resumes [Fig 6. (iii)][61-64]. The SS is then cleaved off by the signal peptidase, which is a membrane-bound protease located on the ER lumen side [Fig 6A.]. While this typical pathway applies to HA, the TMDs of NA and M2 are not removed and are integrated into the ER membrane through the Sec61 translocon [Fig. 6B, C] [65].

Ribosome Golgi A(n) ss (i) SRP

(ii) SR (iii) (iv) NA Translocon HA ER M2

A(n) IAV mRNA Nucleus Cell progeny vRNP

A. Type I (HA) B. Type II (NA) C. Type III (M2) C C C Cytoplasm N N

s SPase ER lumen N ER N ER NN N N C Figure 6. HA, NA and, M2 synthesis, maturation and viral assembly. (i) A newly synthesized N-terminal signal sequence (SS) is recognized and bound by SRP to form the SRP-ribosome-nascent chain complex. (ii) SRP guides the complex to the translocon located on the ER membrane and associates with the SRP receptor (SR). (iii) SRP is released from the ribosome and translation resumes. The protein maturation process is different for the three influenza surface proteins. (Box A) The SS of HA is cleaved off by signal peptidase (SPase) after translocation into the ER, and a C-terminal transmembrane domain integrates HA into the membrane. (Box B) The N-terminal transmembrane domain translocates NA into the ER membrane and inverts during protein synthesis, which positions the NA C-terminal in the ER lumen. (Box C) M2 also utilizes its N-terminal transmembrane domain for ER-targeting, but does not invert during protein synthesis. (iv) After folding and oligomerization, HA, NA and M2 are trafficked to the plasma membrane through the Golgi and are assembled into progeny viruses together with the vRNPs.

Transmembrane insertion into the ER membrane

The Sec61 translocon is a protein-conducting channel that allows nascent polypeptide chains to cross or insert into the membrane depending on the hydrophobicity of the polypeptide region in the channel [66]. During the co-

translational translocation process the hydrophilic polypeptide stretches trav- erse the translocon and enter the ER lumen, whereas the hydrophobic polypeptide stretches are sensed by the lateral gate of the translocon, which then opens to enable these regions to partition into the lipid bilayer [67]. Whether or not the region of the polypeptide is capable of opening the lateral gate can be predicted using the ‘biological’ hydrophobicity scale, DGapp, which calcu- lates the apparent free energy of membrane insertion for linear sequences of amino acids ranging from 19 - 25 [68]. If the prediction for DGapp is < 0 kcal/mol, the linear stretch of amino acids is considered to be hydrophobic and therefore predicted to behave as a TMD and favor integration.

About 25% of known TMDs have DGapp > 0 kcal/mol, which are considered as un-favorable for membrane insertion [69, 70]. These TMDs are mainly part of multi-spanning membrane proteins, which rely largely on the interactions between TMDs for proper insertion [71]. As single-spanning membrane proteins cannot utilize such TMD interactions, very few bitopic proteins have a DGapp > 0 kcal/mol for their single TMD. The predicted DGapp of the TMDs for HA and M2 from the numerous IAV sequences is < 0 kcal/mol, which deems them favored for lateral gate partitioning. However, many subtypes of

NA have a marginally hydrophobic TMD with DGapp > 0 kcal/mol, which are not predicted to favor membrane integration. Insertion of these non-ideal single TMDs can depend on both length [72] and the sequence composition of their C-terminal domain [73]. Kida et al. has found that the marginally hydrophobic TMD can be retained at the translocon and gradually move towards the membrane in a hydrophobicity-dependent manner [74]. The stalling of marginally hydrophobic TMD and the gradual movement as well as the inversion process benefits from the co-translational insertion as the polypeptide remains attached to the ribosome and newly translated polypeptide likely applies a pushing force to facilitate inversion [72].

The orientation of the TMD dictates where the N-terminal and C-terminal domains will be located. The N-terminus of a protein always enters the translocon first and whether the TMD inverts or not can be influenced by positively charged residues that are located next to the TMD. As positive charges are favored to be at the cytoplasmic side of the lipid, TMDs generally invert in the translocon when positive charged residues are located at the N-terminal side [75]. Based on the TMD orientation, the IAV membrane proteins can be

separated into three groups. HA, which utilizes its N-terminal SS for ER targeting is classified as a type I membrane protein since its C-terminal TMD has a Nout-Cin topology [Fig. 6A]. NA, which utilizes its N-terminal TMD for ER targeting is classified as a type II membrane protein since its TMD inverts and resulting a Nin-Cout topology [Fig. 6B]. M2, which also utilizes its N-terminal

TMD is classified as a type III membrane protein as it has a Nout-Cin topology [Fig. 6C].

Protein maturation within the ER lumen

Polypeptide chains fold co-translationally once they leave the ribosomal exit tunnel. In an aqueous environment, proteins generally reach their native state through hydrophobicity collapse. However, the aqueous environment of the cell is a dense milieu with high protein concentration and many of these proteins do not fold sequentially, which creates challenges for folding mediated by hydrophobic collapse. To overcome these problems, cells also possess different types of molecular chaperones, which can slow down the protein folding process by associating with specific regions to prevent aggregation before the distal folding partner is available.

The classical chaperones, which consist of many family members of the heat shock proteins such as Hsp70 and Hsp90, can be found in almost all cellular locations, including the ER. These chaperones bind directly to exposed hydrophobic regions of the polypeptide chain to prevent deleterious side reac- tions during the vulnerable portion of the folding reaction. The lectin chaperones are specific to the ER lumen and these bind to the N-linked glycan mod- ifications that are co-translationally added to secretory proteins by the OST during translocation into the ER lumen [76]. The glycan, which is attached to the Asn residue in the sequence Asn-X-Ser/Thr (where X can be any amino acid except proline), only becomes a substrate for the lectin chaperones once it has been trimmed by glucosidases I and II into a monoglucosylated state [77-79]. In mammalian cells, there are two types of lectin chaperones: calnexin and calreticulin. Calnexin is a membrane protein that mainly binds to glycans found in membrane proximal domains, whereas calreticulin is a soluble protein that associates with glycans that are located deeper into the ER lumen.

Lectin chaperones not only aid in protein folding but are also involved in quality control. When the protein reaches its native conformation, the last glucose on the monoglucosylated side chain is removed by glucosidase II. The protein then dissociates from the lectin chaperones and is exported from the ER for further maturation. If the protein looses its last glucose without reaching its native form, the enzyme glycoprotein glucosyltransferase recognizes it and transfers a glucose on it using UDP-glucose as source. Reglucosylation results in rebinding to calnexin and calreticulin and proceed protein folding [76].

In addition to the lectin chaperones, the ER lumen also contains oxidore- ductases that can accelerate the secretory protein folding rate by catalyzing the proper formation of disulfide bonds. The most abundant oxidoreductase is protein disulfide isomerase (PDI) [80]. One family member of PDI, ERp57, binds to calnexin and calreticulin, which creates a bimolecular pair with chaperone foldase function [76]. This organization accounts for the inability of ERp57 to discern native from nonnative disulfide bonds by relying on the folding recognition by the lectin chaperones.

HA and NA receive several N-linked glycans from the OST during translation, thus it is not surprising that nascent chains for both of these viral proteins have been shown to interact with the lectin chaperones [81, 82]. During synthesis, HA and NA first associate with the membrane bound lectin chaperone calnexin followed by calreticulin, which associates with the regions of the polypeptide that extend into the ER lumen. In HA, each of the Cys residues is located close to an N-linked glycan, suggesting calnexin or calreticulin associated with ERp57 to guide the proper formation of disulfide bound. This organization is especially important for the formation of the large disulfide loop that links a region in HA1 with a distal region in HA2 [83]. For NA, the glycans on the head domain have been shown to be essential for folding, whereas the ones on the stalk region are dispensable, which indicates that the formation of the intramolecular disulfide bonds in the head domain likely requires the lectin mediated delivery of ERp57. In addition, NA has also been shown to transiently associate with the classical Hsp70 chaperone BiP, but the requirement for this interaction remains unclear [84].

For both NA and HA, oligomerization is the last step in their maturation process, however the process by which this occurs is very different between these two proteins. After the individual HA monomers are synthesized and

folded, HA starts to form homo-trimers in the ER [85, 86]. In contrast, the oligmerization of NA occurs in a two-step process, where NA dimerizes co- translationally and the dimers assemble into tetramers post-translationally. For the co-translational dimerization of NA it was shown that the Cys residues in the stalk region form intermolecular disulfide bonds even before the complete synthesis of the second monomer [82]. This finding supports previous data by Saito et al. where epitope specific NA monoclonal antibodies were used to show that NA dimerization happens long before tetramer formation [87]. In line with the earlier work by Wang et al, da Silva and Nordholm showed that oligomerization of the NA TMD is necessary for the optimal assembly of the tetrameric head domain [88]. They further demonstrated that the NA TMD is an amphipathic helix and that the polar residues are the main driving force of its oligomerization [89]. Although it has not been shown directly, it is possible that the polar regions of the TMD dimer pairs undergo a slight shift during the tetramerization process that supports that assembly of the head domain.

Structure and function of NA

Structure of NA

NA is a tetrameric protein and each monomer is composed of four domains: a short cytosolic N-terminal domain, a TMD, a stalk region and a globular enzymatic head domain. The globular enzymatic head domain possesses a 6- bladed propeller shape, and each of the blades is formed by four antiparallel b-sheets that are connected by loops and stabilized by intermolecular disulfide bonds and connected by loops. At the center of each globular head domain is a deep cavity that functions as the enzymatic pocket. The enzymatic pocket of NA is composed of functional amino acids, which bind the sialic acid backbone, and the catalytic residue Tyr 402 [Fig 7. upper right monomer]. By binding to the sialic acid backbone, the functional residues trigger conformational change in sialic acid that makes it more susceptible to nucleophilic attack from Tyr 402. Depending on the strain and subtype, one to two Ca2+ ions have been resolved close to the catalytic site in each head domain monomer, and some structures have also resolved a central Ca2+ ion that interacts with all four monomers [90] [Fig 7.]. Even though NA contains an enzymatic pocket in each of the monomers, it requires its quartenary tetrameric structure to be en- zymatically active. Currently, it is unclear why IAV NA has evolved to function as a tetramer, but it it has been suggested that the active state requires inter-subunit interactions, such as interaction with the central Ca2+ [91], the formation of salt bridges [92].

Figure 7. Crystal structure of a subtype 1 influenza NA head domain. Four identical monomers compose the NA head, each carrying an enzymatic pocket. The NA inhibitor Zanamivir (red) is shown bound within the enzymatic pocket. In the upper right monomer (yellow), the enzymatic pocket is highlighted in sticks (blue). (Functional amino acid: blue; active site Tyr 402: green) Also shown are the Ca2+ ions (cyan), two in each monomer and one in the center that interacts with all four monomers. (PDB: 3TI5) [93]

Function of NA

Once NA has properly assembled into a tetramer it is able to function as a sialidase, an enzyme which cleaves terminal sialic acid residues from oligosaccharide chains. When NA encounters a sialylated glycoconjugate, the terminal sialic acid residue binds deep in the catalytic pocket of NA and the first few following oligosaccharides make small, but important interactions with residues that form the cavity. The strong ionic interactions between several conserved changed amino acids (Arg118, Asp151, Arg152, Arg225, Glu277, Arg293, Arg368 and Tyr402) and the sialic acid activate the C2 atom, trigger- ing a conformational change from chair to boat (the transition state). The conformation change makes the C2 atom more susceptible for nucleophilic at- tacking by the deprotonated hydroxyl group of Tyr 402 presumably due to reposition the ring structure. Following the attack, Tyr 402 remains covalently linked to the sialic acid C2 atom and the underlying oligosaccharide (HOR) is released. The hydrolysis reaction is completed by a water molecule, which is deprotonated by general base catalysis, generating a nucleophilic hydroxide ion. The hydroxide ion attacks and breaks the ester bond between C2 atom of

sialic acid and the Tyr, recharging the Tyr with a proton to recreate its natural hydroxylated side chain [Fig 8.] [94].

Tyr 402 Tyr 402 Tyr 402 HOR OH O- Activate HO OH O O - HO OH O substrate HO OH OR O O O HO HO 6 O 2 OR O - 4 HO 6 2 HN O 5 3 5 4 3 O- HN HN OH O O O Enzyme - substrate Substrate Binding Substrate transition state intermediate

Tyr 402 Tyr 402 H H O - OH HO OH O O HO OH O O O HO 6 O 2 OH HO 4 - 5 3 HN O HN OH O O Sialic acid release Enzyme recharge Figure 8. Mechanism of sialic acid cleavage by NA. Upon binding to the NA enzymatic pocket, the sialic acid C2 atom is activated by a conformational change (from chair to boat) of the 6-atom ring structure. The deprotonated hydroxyl group of Tyr 402 then performs a nucleophilic attack on the activated C2 atom, creating the enzyme-substrate intermediate and releasing the oligosaccharide side chains. Deproto- nated water molecule nucleophilic attack the ester bond between Tyr 402 and the C2 atom, releasing the sialic acid and recharge Tyr 402 with a proton. This illustration is modified from Shtyrya, 2009

During an IAV infection cycle, the enzymatic function of NA plays many important roles in several different viral processes. The upper respiratory epithelium is protected by a layer of mucus, which contains a large portion of heavily glycosylated proteins such as mucins. In order to infect epithelial cells, IAVs first need to pass through the mucus layer. The IAV surface protein HA, however, can bind to mucins, which prevents the virus from penetrating the mucus layer [95]. By cleaving off sialic acid residues on proteins such as mucins, NA helps the virus to penetrate the mucus layer and reach the epithelial cells where it can initiate a viral infection [96, 97] [Fig. 9(i)]. After IAVs bud off from the host cell, HA-mediated receptor binding can keep the virus attached to the cell surface. NA facilitates the release of these newly formed

viral particles by removing sialic acid residues from the infected host cell surface [98, 99][Fig. 9(ii)]. Viruses can also associate with themselves due to HA associations with the glycoconjugates on HA and NA from neighboring viruses. NA can separate these agglutinated viruses by cleaving off the sialic acids on the N-linked glycans present on the HA and NA molecules that are located on the viral surface [100] [Fig. 9(iii)]. This function may explain the relationship between NA activity and IAV transmissibility [101, 102], as single viruses are more likely to be transmitted in small aerosol droplets.

Sialylated Glycan Galactose

SA (iii)

+ H2 O HA NA SA hydrolysis IAV IAV IAV

IAV (ii) Mucin (i)

Ciliated Mucin Infected cell epithelial secreted cell cell

Figure 9. NA sialidase activity contributes to viral entry and release. (i) In the respiratory tract, IAVs have to penetrate the protective mucus layer, rich in the sialylated mucin glycoproteins, to allow for infection. NA cleaves off the sialic acid (SA) from mucin and facilitates movement through the mucus layer. (ii) Following viral budding, the virus can remain attached to the cell surface via HA-sialylated receptor binding. NA promotes viral release by removing sialic acid from the receptors. (iii) Both NA and HA are glycoproteins, which lead to HA-mediated virus aggregation. NA cleaves off sialic acid and separates the viral particles.

Due to the essential role of NA activity in a viral infection process and its rigid and conserved catalytic site, it has been a main target for anti-influenza drugs. The currently FDA-approved drugs include TamifluTM (oseltamivir), and RelenzaTM (zanamivir) [Fig 10.], and peramivir (Rapivab) [103, 104]. These drugs are designed to mimic the structure of the sialic acid transition state, thereby competing with the natural substrate to inhibit the enzymatic activity of NA. However, the treatment with these inhibitors is less than ideal as the treatment window is 48 h after the onset of symptoms and viruses have

been shown to rapidly develop mutations that decrease the binding affinity of the inhibitors [1].

Figure 10. Structures of sialic acid, Zanamivir and Oseltamivir. Zanamivir and Osel- tamivir are current FDA-approved synthetic molecules that mimic the transition state of sialic acid binding to the NA enzymatic pocket. The blue portions highlight the parts oft he inhibitors that are identical to sialic acid.

Antibodies against NA

When IAVs initiate an infection, the host immune system aims to defend against it and limit the spread of the virus. In humans, the response involves both the innate and adaptive immune system. The innate immune system is the first to sense the invasion of the foreign pathogen and it assists in the acti- vation of the adaptive immune system, which ultimately results in the produc- tion of antibodies against the IAV antigens.

During a natural IAV infection, humans have been shown to generate antibodies against several different viral proteins. The majority of these antibodies target the surface glycoproteins HA and NA, which are some of the more abundant viral proteins that are expressed. Chen et al. have examined the plasma blasts from 16 IAV infected patients and found that 1/3 of these cells produce antibodies that react with HA, 1/4 produce antibodies against NA, and the remaining fraction produced antibodies that reacted with other viral proteins [105].

NA antibodies have been shown to protect against IAV infection in humans back in the 1970’s [106, 107]. More recent clinical trials have reexamined

these findings and demonstrated that influenza virus protection correlates to both anti-NA and anti-HA serum antibody titers [108, 109]. Experiments per- formed in animal models have additionally shown that a strong induction of NA-specific antibodies provides protection against a lethal virus challenge [105, 110].

Antibodies generally have an effector function in recruiting immune cells to eliminate infections. One of the mechanisms is known as antibody-dependent cell-mediated cytotoxicity (ADCC), where NK cells recognize and bind to the Fc portion of the antibody that is associated with an antigen on the infected cell surface and lyse it. An additional clearance process is known as antibody- dependent phagocytosis (ADP), where macrophages are recruited to phagocy- tose antibody-coated viral particles or apoptosis cells. Even though recent studies have indicated anti-NA antibodies have ADCC effect in vitro, the primary effector function of NA antibodies in humans is still unclear [111].

Similar to HA antibodies that can neutralize viral particles by inhibiting its binding function, NA antibodies can also have a neutralizing effect on its enzymatic function. Although NA antibodies have not been studied in great de- tail, they can be separated into the following three categories based on their ability to neutralize NA activity: -i- NA activity inhibiting (NAI) antibody, which generally bind the enzymatic pocket and block the accessibility of the sialylated glycoconjugate; -ii- Partial NA activity inhibiting (semi-NAI), which bind near the enzymatic pocket and interfere with sialylated glycoconjugate binding by steric hinderance [112]; and -iii- Non NA activity inhibiting (non-NAI), which bind to NA but do not affect its enzymatic function [113].

Currently it is considered that the neutralization effect directly correlates to the NAI activity of these antibodies, making them similar to NA inhibitors. Supporting this correlation, in vitro studies have shown that both NAI and semi-NAI antibodies can reduce viral propagation in tissue culture [112] and in an ex vivo upper respiratory epithelial cell systems containing a mucus layer [96]. However, it is not clear if the effectiveness between these two systems is equal.

Generally, antibodies produced by plasma cells have exactly the same antigen recognition as the membrane-bound Ig receptor on its precursor B lym- phocytes. Recent work has shown that NA antibodies can cross-react within the same subtype for both N1 and N2 [105, 114] raising the question whether

NA-specific antibodies can provide a broad protection effect against virus across different NAs within the same subtype. Ferret and mouse model experiments have shown that immunization with NA antigen not only can protect from virus with matched IAV strain but also protect from IAV carrying NA from a different strain but the same subtype [115, 116]. The cross protection is likely due to the contribution of conserved epitopes that gives cross-reactiv- ity between the different NAs [113].

Results summary

Paper I. Polar Residues and Their Positional Context Dictate the Trans-membrane Domain Interactions of Influenza A Neuraminidases.

NA is a tetrameric membrane protein that is comprised of a head domain, a stalk linker region and a TMD. In the cell, the assembly of subtype 1 NA is a cooperative proceess that involves oligomerization of both the enzymatic head domain and the TMD. Here, we show that the TMDs from all NA subtypes except for N2 are amphipathic a-helices that contain a polar face, which mediate their homo-oligomerization. Across the different subtypes, the strength of the TMD interactions inversely correlates with the TMD hydrophobicity, indicating that polar residues are responsible for the interaction. A temporal analysis of the NA TMD from human H1N1 strains indicates that hydrophilic drift is occurring in the TMD that leads to a stronger interaction in TMD overtime, suggesting this non exposed region of NA may be under selection pressure.

Paper II. Type II Transmembrane Domain Hydrophobicity Dictates the Co- translational Dependence for Inversion.

Single-pass membrane proteins have one chance to insert into the ER membrane during synthesis. Therefore, the TMDs from this class of proteins are generally of sufficient hydrophobicity to mediate their insertion into the membrane. However, in human H1N1 viruses the NA TMD shows evidence of amphiphilic drift with the more recent NAs encoding for a marginally hydrophobic TMD. By analyzing a series of increasing length NAs, we found that the correct membrane insertion of these marginally hydrophobic TMDs is related to the arrangement of NA’s domains. During protein translation, these N-terminal TMDs are hydrophobic enough to guide the ribosome-nascent chain complex to the translocon. However, the translation of the long NA C- terminus is necessary for the marginally hydrophobic TMDs to invert and ori-

entate in the proper topology for insertion into the lipid membrane. Quantifi- cation of our results showed that the efficiency of NA insertion and inversion reaches 50% when the polypeptide length following the TMD is approxi- mately 100 amino acids. This length is consistent with that of the domains which follow all of the predicted type II human membrane proteins that possess a marginally hydrophobic TMD. Together, this study shows how the co- translational translocation process contributes to the lipid bilayer integration of single-pass membrane protein with a marginally hydrophobic TMD.

Paper III. Structural Restrictions for Influenza Neuraminidase Activity Pro- mote Adaptation and Diversification.

Although all IAVs contain NA, significant variation exists in the NAs that circulate in nature. Even within the same subtype, NAs show variation in their properties, such as substrate binding affinity, enzyme activity, and protein thermo- and pH- stability. In this study, we investigated what is responsible for the property variation of subtype 1 NAs (N1) in human IAVs. We first determined that the central calcium ion, which associates with all four monomers, is a major stability factor for NA. By selectively substituting the amino acids that alter the structure of the central calcium binding pocket, the affinity of NA for this calcium ion changes, and subsequently stability of NA changes. The flexibility in the structure that enables these changes at the interface were also found to allow different NA monomers to assemble together into a tetrat- mer, providing a mechanism to further increase the diversity of NA. These results indicate that the structural requirements for NA function provides the enzyme with the advantages of increased mutational tolerance and access to more diversity in its properties.

Paper IV. Analysis of IAV Replication and Co-infection Dynamics by a Ver- satile RNA Viral Genome Labeling Method.

Delivering genetic material to the proper cellular compartment for replication is the ultimate goal for all viruses. The segmented nature of the IAV genome complicates this process, but it also allows IAVs to reassort genome segments from different viruses during a co-infection. In this paper, we developed an in situ RNA labeling approach to visualize the IAV gene segments in cells during an infection. The high specificity of the labeling allowed us to track IAV gene segment trafficking from cell entry through nuclear export. The results

showed that the half time for a single viral particle enter the cell is ~5 min and that the nuclear import of the gene segments is an efficient process. We used the quantitative and spatial information to define the IAV infection stage in single cells and to investigate the temporal requirements for IAV reassortment. By taking advantage of the single nucleotide specificity, it was possible to track the gene segments of two isogenic virus during co-infection experiments. Our results show that the ability of a second virus to replicate in the same cell is limited once the first virus has reached its replication stage [Fig 11.]. This temporal restriction combined with the spatial requirement for a cell co-infection are likely contributing factors to the low prevalence of reassortment events in nature.

Figure 11. Model of Cell susceptibility to secondary infection. Over one course of infection, five infection stages were defined based on the spatial and quantity information acquried through gene segment labeling. The proportions of cell population from each stage were plot against the corresponding infection time from a single infection. The time window that whether the cells are susceptible to co-infection are shown above the graph. Data from Paper IV.

Conclusions and future perspective

IAVs commonly circulate in the human population and are a major contributing factor to the yearly influenza epidemics and are responsible for the oc- casional influenza pandemic outbreaks. The severity of this disease is associated with the ability of IAVs to quickly evolve, which results in the mismatch between pre-existing immunity of each individual and current circulating IAV strains. IAVs evolve through two primary mechanisms: antigenic drift and antigenic shift. In this thesis work, I have focused on studying how the NA surface antigen from H1N1 IAVs has evolved with respect to the limitations imposed by the host cellular machinery and its functional requirements.

Much of the work presented here focused on analyzing non-exposed amino acids of NA that reside in the viral envelope, which is quite different from the common approach of analyzing amino acids on the NA ectodomain that contains the enzymatic region. The rationale behind this comes from previous work from our lab that has shown that the TMD of NA is not only a membrane anchor but also aids in its tetramerization [88]. Oligomerization of TMDs is driven by polar-polar interaction as each TMD is an amphipathic a-helix. Over time, the NA TMD in human H1N1 viruses has become less and less hydrophobic, which provides a stronger interaction in TMD among monomers (Paper I). However, the hydrophilic drift goes against the cellular hydrophobicity requirement of ER insertion of single TMD protein. This raises the question of how the hydrophilic drift is possible to occur, as the IAV protein synthesis process is fully dependent on the cellular machinery. In paper II, we found that the key reason why NA can tolerate having a marginally hydrophobic TMD is due to the location of its TMD on the N-terminus followed by a long C-terminal domain. This enables the co-translational synthesis process to aid in the positioning and inversion of these TMDs for proper insertion into the membrane.

Paper II provided a comprehensive analysis on the relationship between TMD hydrophobicity and how NA as well as cellular proteins with this characteristic are able to integrate into the ER membrane. Specifically, we showed

that proteins with marginally TMDs require the co-translational translocation process to facilitate their membrane integration. Potentially this requirement is related to the need for the amphipathic helix to adjust its orientation so that the hydrophobic side faces the lateral gate of the Sec translocon. In this manner, the co-translational process would provide the additional time by keeping the nascent chain attached to the ribosome. When we considered that the ribosome exit tunnel holds ~40 aa and the Sec translocon holds ~20 aa, our results showing that the C-terminal length needs to be at least 70 aa to initiate inversion would support such a conclusion as the extra 10 aa would provide slack in the polypeptide chain and about 2 seconds of additional time for the inversion and positioning to occur. We demonstrated this more directly by showing that another TMD could be inverted by elongating the C-terminal domain (Pa- per II). However, it is still not certain how the additional polypeptide length and time contribute to inversion process as it is possible that this requirement is related to the binding of chaperones from within the ER lumen. In the future, it will be interesting to investigate whether hydrophobic and marginally hydrophobic TMDs use the same translocation machinery.

Although the results in previous work and paper I show that the TMD contributes to the tetrameric assembly of the NA head domain, we still do not know why. One interesting observation we made was that the TMD of NA is connected to the stalk, which feeds into the base of each head domain around the axis of symmetry. More recent NA structures have shown that directly above this region lies a calcium binding pocket, which must be occupied for NA to function. This pocket has a low affinity for calcium and has previously been suggested to aid in NA stability. The reason that NA has evolved to require calcium binding in such a low affinity pocket is probably due to calcium being a unique factor that differs in concentration among different environ- ments. Therefore, changes in calcium ion concentration can alter NA enzymatic properties, which could be a simple mechanism for regulating NA function (Paper III).

Besides changing NA gradually, IAVs can also evolve through antigenic shift where NAs from multiple viruses exchange during the co-infection process, resulting in a new virus strain. The approach I developed during my thesis work for detecting the cellular localization of the RNA genomic segments should make it possible to examine numerous details of the IAV genome in cells that were not previously possible. The method provides the ability to label all eight short IAV RNA gene segments with high specificity, enabling us

to visualize the delivery of the IAV genome to the nucleus for the first time (Paper IV). In contrast to protein labeling, this method can easily differentiate between very similar viruses. By combining IAV gene segment localization profile with semi-quantitative information of each genome at a single-cell level, the infection stage of each individual cell can be identified among a large population. This provides a new way of identifying the infection stages across a population of cells. We took advantage of the single-nucleotide specificity to study the temporal requirement of productive co-infection in a non- biased setting. The results from this analysis showed that the replication of a second IAV genome is limited when the first IAV genome is towards the end of the replication cycle, likely explaining the sporadic event of antigenic shift. However, the mechanism that limits the nuclear import of the second viral genome remains unknown. Two possibilities for this observation are potentially an active blockage process of IAV or the lack of certain components of the cellular machinery.

Taken together, the studies that I have been involved with have provided new information for how NA has evolved as a protein and fulfilled the requirements imposed by the cellular machinery that are necessary for its synthesis, assembly and trafficking. These findings offer potential insight into which di- rection NA can or cannot evolve that may provide some guidance with respect to predicting NA evolution and future vaccine strains. The labeling methodol- ogy I helped to develop opens up the ability to examine aspects of the infection process that were not previously possible, such as how the cellular machinery facilitates or restricts the viral genome from entering the cell, trafficking between organelles and importing into the nucleus. In the future, it will be interesting to investigate the genome trafficking process in a variety of cell types including airway epithelial cells and immune cells both in vitro and in vivo.

Sammanfattning på svenska

När influensavirus infekterar människor kan det orsaka mild till akut luftvägs- infektion. Förutom individens existerande immunitet och hälsostatus, är variation i viruset också en viktig faktor för utfallets allvarlighet. Hos viruset är det ofta relaterat till dess evolutionsförmåga att undvika individens existerande immunitet. Mekanismen för detta är att variera proteinerna på virusets yta, hemagglutinin (HA) och neuraminidas (NA). Detta sker på två sätt, med hjälp av antigeniskt drift (ackumulering av mutationer i dessa proteiner), eller antigeniskt skift (utbyte av HA eller NA mellan två olika influensavirus som infekterar samma cell).

I mitt arbete har jag använt NA för att förstå evolutionen hos influensaviruset under påtryckning av immunförsvaret, hur den samtidigt kan bevara nödvän- diga proteinfunktioner, samt hur den samtidigt kan anpassa sig för att utnyttja den infekterade cellen för att göra fler viruskopior.

NA är ett svampformat membranprotein, vars ”huvud” (kallad huvuddomän) är lokaliserad utanför cellen, och vars ”fot” (kallad transmembran domän) är inuti cellmembranet. Vanligtvis sker antigeniskt drift på ytan av proteiner, eftersom det är där antikroppar har möjlighet att binda. Överraskande har vi i mitt arbete hittat två gömda proteinregioner som också genomgår antigeniskt drift. Den ena är i membranbundna transmembrana domänen, och den andra är i centrala fickan på huvuddomänen som calciumjoner vanligtvis binder till. Denna upptäckt indikerar att NA, som en helhet istället för som separata do- mäner, evolverar för att förhindra antikroppsigenkänning.

För att studera antigeniskt skift har jag utvecklat en väldigt känslig och specifik metod för RNA-detektion för att visualisera influensagenomet under in- fektionsprocessen, som gör det möjligt att i detalj studera vart genomet befin- ner sig i varje stadie. Och eftersom metoden är så specifik går det att studera utbyte av gensegment mellan olika virus utan att behöva titta på proteiner.

Acknowledgments

First, I would like to thank my supervisor, Rob Daniels, for giving me opportunity to work in your lab. During these years, you have shown me the fun working in research. Thank you for being so helpful and always providing so many ideas for the projects. I have learned a lot and I am very grateful for the experience.

A special thanks to Jan-Willem de Gier for being my co-supervisor and to Stefan Nordlund, Pia Ädelroth and Martin Ott for organizing the phD program and for doing the check points during my phD time.

Thanks to Professor Schwemmle, Professor Masucci, Patrik Ellström and Professor Högbom for being my opponent and committee.

Thanks to the former and current lab members: Diogo, you were my ‘guide man’ in the lab and I am so grateful to have you as my teacher. You are always so supportive and taking care of me. Johan, discussing with you always in- spires me, and it has been a quite year without you. Also, thanks for fixing all my Mac problems. Hao, 感谢你在工作中和生活中给我的一切支持 ❤. An- nika, thank you for go through my thesis and give me advice on the thesis manuscript. Henrik, thank you for sharing your opinions about Swedish soci- ety and Swedish education. Rebecca, it has been nice working with you.

Thanks to my collaborators: Thanks to Mats Nilsson for your valuable input in the project and give me opportunity to be in your lab. Ivan, you are such a nice guy and always support me when I need help with experiment. Thanks for the hard work and expertise contributed in the padlock project. Xiaoyan, thanks for helping me with microscope and imaging analysis.

Thanks to the everyone in DBB for creating a convenient working environment. And a big hug to my previous and current lab neighbors at DBB: Cata, Beta, Pedro, Bill, Patrick, Thomas, Zhe, Grant, Nir, Candan, Chenge,

Weihua, Biao, Xin, Fan, Huabin, thank you for the accompany and for making these years so enjoyable.

At last, thanks to my parents, my parents-in-law, and my uncle and aunt (我的姨和舅舅) for supporting me during these years. And also to my little family: 你们是我前行的动力和方向.

References

1. Paules, C. and K. Subbarao, Influenza. Lancet, 2017. 390(10095): p. 697-708. 2. Dou, D., et al., Influenza A Virus Cell Entry, Replication, Virion Assembly and Movement. Front Immunol, 2018. 9: p. 1581. 3. McGeoch, D., P. Fellner, and C. Newton, Influenza virus genome consists of eight distinct RNA species. Proc Natl Acad Sci U S A, 1976. 73(9): p. 3045-9. 4. Harris, A., et al., Influenza virus pleiomorphy characterized by cryoelectron tomography. Proc Natl Acad Sci U S A, 2006. 103(50): p. 19123-7. 5. Gamblin, S.J. and J.J. Skehel, Influenza hemagglutinin and neuraminidase membrane glycoproteins. J Biol Chem, 2010. 285(37): p. 28403-9. 6. Getie-Kebtie, M., et al., Label-free mass spectrometry-based quantification of hemagglutinin and neuraminidase in influenza virus preparations and vaccines. Influenza Other Respir Viruses, 2013. 7(4): p. 521-30. 7. Arranz, R., et al., The structure of native influenza virion ribonucleoproteins. Science, 2012. 338(6114): p. 1634-7. 8. Moeller, A., et al., Organization of the influenza virus replication machinery. Science, 2012. 338(6114): p. 1631-4. 9. Pflug, A., et al., Structure of influenza A polymerase bound to the viral RNA promoter. Nature, 2014. 516(7531): p. 355-60. 10. Fodor, E., B.L. Seong, and G.G. Brownlee, Photochemical cross- linking of influenza A polymerase to its virion RNA promoter defines a polymerase binding site at residues 9 to 12 of the promoter. J Gen Virol, 1993. 74 ( Pt 7): p. 1327-33. 11. Yoon, S.W., R.J. Webby, and R.G. Webster, Evolution and ecology of influenza A viruses. Curr Top Microbiol Immunol, 2014. 385: p. 359-75. 12. Morens, D.M., J.K. Taubenberger, and A.S. Fauci, The persistent legacy of the 1918 influenza virus. N Engl J Med, 2009. 361(3): p. 225-9.

13. Parvin, J.D., et al., Measurement of the mutation rates of animal viruses: influenza A virus and poliovirus type 1. J Virol, 1986. 59(2): p. 377-83. 14. Suarez, P., J. Valcarcel, and J. Ortin, Heterogeneity of the mutation rates of influenza A viruses: isolation of mutator mutants. J Virol, 1992. 66(4): p. 2491-4. 15. Suarez-Lopez, P. and J. Ortin, An estimation of the nucleotide substitution rate at defined positions in the influenza virus haemagglutinin gene. J Gen Virol, 1994. 75 ( Pt 2): p. 389-93. 16. Bloom, J.D., An experimentally determined evolutionary model dramatically improves phylogenetic fit. Mol Biol Evol, 2014. 31(8): p. 1956-78. 17. Pauly, M.D., M.C. Procario, and A.S. Lauring, A novel twelve class fluctuation test reveals higher than expected mutation rates for influenza A viruses. Elife, 2017. 6. 18. Rambaut, A., et al., The genomic and epidemiological dynamics of human influenza A virus. Nature, 2008. 453(7195): p. 615-9. 19. Sobel Leonard, A., et al., The effective rate of influenza reassortment is limited during human infection. PLoS Pathog, 2017. 13(2): p. e1006203. 20. Lowen, A.C., Constraints, Drivers, and Implications of Influenza A Virus Reassortment. Annu Rev Virol, 2017. 4(1): p. 105-121. 21. Bottcher-Friebertshauser, E., et al., The hemagglutinin: a determinant of pathogenicity. Curr Top Microbiol Immunol, 2014. 385: p. 3-34. 22. Weis, W., et al., Structure of the influenza virus haemagglutinin complexed with its receptor, sialic acid. Nature, 1988. 333(6172): p. 426-31. 23. Rust, M.J., et al., Assembly of endocytic machinery around individual influenza viruses during viral entry. Nat Struct Mol Biol, 2004. 11(6): p. 567-73. 24. Chen, C. and X. Zhuang, Epsin 1 is a cargo-specific adaptor for the clathrin-mediated endocytosis of the influenza virus. Proc Natl Acad Sci U S A, 2008. 105(33): p. 11790-5. 25. de Vries, E., et al., Dissection of the influenza A virus endocytic routes reveals macropinocytosis as an alternative entry pathway. PLoS Pathog, 2011. 7(3): p. e1001329. 26. Guinea, R. and L. Carrasco, Requirement for vacuolar proton-ATPase activity during entry of influenza virus into cells. J Virol, 1995. 69(4): p. 2306-12. 27. Pinto, L.H. and R.A. Lamb, The M2 proton channels of influenza A and B viruses. J Biol Chem, 2006. 281(14): p. 8997-9000. 28. Yoshimura, A. and S. Ohnishi, Uncoating of influenza virus in endosomes. J Virol, 1984. 51(2): p. 497-504. 29. Bullough, P.A., et al., Structure of influenza haemagglutinin at the pH of membrane fusion. Nature, 1994. 371(6492): p. 37-43.

30. White, J.M. and G.R. Whittaker, Fusion of Enveloped Viruses in Endosomes. Traffic, 2016. 17(6): p. 593-614. 31. Beck, M. and E. Hurt, The nuclear pore complex: understanding its function through structural insight. Nat Rev Mol Cell Biol, 2017. 18(2): p. 73-89. 32. Stewart, M., Molecular mechanism of the nuclear protein import cycle. Nat Rev Mol Cell Biol, 2007. 8(3): p. 195-208. 33. Eisfeld, A.J., G. Neumann, and Y. Kawaoka, At the centre: influenza A virus ribonucleoproteins. Nat Rev Microbiol, 2015. 13(1): p. 28-41. 34. Martin, K. and A. Helenius, Transport of incoming influenza virus nucleocapsids into the nucleus. J Virol, 1991. 65(1): p. 232-44. 35. Kalderon, D., et al., A short amino acid sequence able to specify nuclear location. Cell, 1984. 39(3 Pt 2): p. 499-509. 36. Wang, P., P. Palese, and R.E. O'Neill, The NPI-1/NPI-3 (karyopherin alpha) binding site on the influenza a virus nucleoprotein NP is a nonconventional nuclear localization signal. J Virol, 1997. 71(3): p. 1850-6. 37. Cros, J.F., A. Garcia-Sastre, and P. Palese, An unconventional NLS is critical for the nuclear import of the influenza A virus nucleoprotein and ribonucleoprotein. Traffic, 2005. 6(3): p. 205-13. 38. Wu, W.W., L.L. Weaver, and N. Pante, Ultrastructural analysis of the nuclear localization sequences on influenza A ribonucleoprotein complexes. J Mol Biol, 2007. 374(4): p. 910-6. 39. Plotch, S.J., et al., A unique cap(m7GpppXm)-dependent influenza virion endonuclease cleaves capped RNAs to generate the primers that initiate viral RNA transcription. Cell, 1981. 23(3): p. 847-58. 40. Guilligay, D., et al., The structural basis for cap binding by influenza virus polymerase subunit PB2. Nat Struct Mol Biol, 2008. 15(5): p. 500-6. 41. Dias, A., et al., The cap-snatching endonuclease of influenza virus polymerase resides in the PA subunit. Nature, 2009. 458(7240): p. 914-8. 42. Reich, S., et al., Structural insight into cap-snatching and RNA synthesis by influenza polymerase. Nature, 2014. 516(7531): p. 361- 6. 43. Bier, K., A. York, and E. Fodor, Cellular cap-binding proteins associate with influenza virus mRNAs. J Gen Virol, 2011. 92(Pt 7): p. 1627-34. 44. York, A. and E. Fodor, Biogenesis, assembly, and export of viral messenger ribonucleoproteins in the influenza A virus infected cell. RNA Biol, 2013. 10(8): p. 1274-82. 45. Poon, L.L., et al., Direct evidence that the poly(A) tail of influenza A virus mRNA is synthesized by reiterative copying of a U track in the virion RNA template. J Virol, 1999. 73(4): p. 3473-6.

46. Huet, S., et al., Nuclear import and assembly of influenza A virus RNA polymerase studied in live cells by fluorescence cross-correlation spectroscopy. J Virol, 2010. 84(3): p. 1254-64. 47. York, A., et al., Isolation and characterization of the positive-sense replicative intermediate of a negative-strand RNA virus. Proc Natl Acad Sci U S A, 2013. 110(45): p. E4238-45. 48. Turrell, L., et al., The role and assembly mechanism of nucleoprotein in influenza A virus ribonucleoprotein complexes. Nat Commun, 2013. 4: p. 1591. 49. Ye, Q., R.M. Krug, and Y.J. Tao, The mechanism by which influenza A virus nucleoprotein forms oligomers and binds RNA. Nature, 2006. 444(7122): p. 1078-82. 50. Lee, N., et al., Genome-wide analysis of influenza viral RNA and nucleoprotein association. Nucleic Acids Res, 2017. 45(15): p. 8968- 8977. 51. Williams, G.D., et al., Nucleotide resolution mapping of influenza A virus nucleoprotein-RNA interactions reveals RNA features required for replication. Nat Commun, 2018. 9(1): p. 465. 52. Fodor, E., The RNA polymerase of influenza a virus: mechanisms of viral transcription and replication. Acta Virol, 2013. 57(2): p. 113- 22. 53. Elton, D., et al., Interaction of the influenza virus nucleoprotein with the cellular CRM1-mediated nuclear export pathway. J Virol, 2001. 75(1): p. 408-19. 54. Watanabe, K., et al., Inhibition of nuclear export of ribonucleoprotein complexes of influenza virus by leptomycin B. Virus Res, 2001. 77(1): p. 31-42. 55. Zerial, M. and H. McBride, Rab proteins as membrane organizers. Nat Rev Mol Cell Biol, 2001. 2(2): p. 107-17. 56. Amorim, M.J., et al., A Rab11- and microtubule-dependent mechanism for cytoplasmic transport of influenza A virus viral RNA. J Virol, 2011. 85(9): p. 4143-56. 57. Eisfeld, A.J., et al., RAB11A is essential for transport of the influenza virus genome to the plasma membrane. J Virol, 2011. 85(13): p. 6117- 26. 58. Momose, F., et al., Apical transport of influenza A virus ribonucleoprotein requires Rab11-positive recycling endosome. PLoS One, 2011. 6(6): p. e21123. 59. de Castro Martin, I.F., et al., Influenza virus genome reaches the plasma membrane via a modified endoplasmic reticulum and Rab11- dependent vesicles. Nat Commun, 2017. 8(1): p. 1396. 60. Lehninger, A.L., D.L. Nelson, and M.M. Cox, Lehninger principles of biochemistry. 6th ed. 2013, New York: W.H. Freeman.

61. Shan, S.O., S.L. Schmid, and X. Zhang, Signal recognition particle (SRP) and SRP receptor: a new paradigm for multistate regulatory GTPases. Biochemistry, 2009. 48(29): p. 6696-704. 62. Walter, P. and G. Blobel, Translocation of proteins across the endoplasmic reticulum III. Signal recognition protein (SRP) causes signal sequence-dependent and site-specific arrest of chain elongation that is released by microsomal membranes. J Cell Biol, 1981. 91(2 Pt 1): p. 557-61. 63. Gilmore, R., P. Walter, and G. Blobel, Protein translocation across the endoplasmic reticulum. II. Isolation and characterization of the signal recognition particle receptor. J Cell Biol, 1982. 95(2 Pt 1): p. 470-7. 64. Gorlich, D., et al., A mammalian homolog of SEC61p and SECYp is associated with ribosomes and nascent polypeptides during translocation. Cell, 1992. 71(3): p. 489-503. 65. Walter, P. and A.E. Johnson, Signal sequence recognition and protein targeting to the endoplasmic reticulum membrane. Annu Rev Cell Biol, 1994. 10: p. 87-119. 66. Van den Berg, B., et al., X-ray structure of a protein-conducting channel. Nature, 2004. 427(6969): p. 36-44. 67. Gogala, M., et al., Structures of the Sec61 complex engaged in nascent peptide translocation or membrane insertion. Nature, 2014. 506(7486): p. 107-10. 68. Hessa, T., et al., Molecular code for transmembrane-helix recognition by the Sec61 translocon. Nature, 2007. 450(7172): p. 1026-30. 69. White, S.H. and G. von Heijne, How translocons select transmembrane helices. Annu Rev Biophys, 2008. 37: p. 23-42. 70. Ojemalm, K., et al., Orientational preferences of neighboring helices can drive ER insertion of a marginally hydrophobic transmembrane helix. Mol Cell, 2012. 45(4): p. 529-40. 71. De Marothy, M.T. and A. Elofsson, Marginally hydrophobic transmembrane alpha-helices shaping membrane protein folding. Protein Sci, 2015. 24(7): p. 1057-74. 72. Dou, D., et al., Type II transmembrane domain hydrophobicity dictates the cotranslational dependence for inversion. Mol Biol Cell, 2014. 25(21): p. 3363-74. 73. Junne, T. and M. Spiess, Integration of transmembrane domains is regulated by their downstream sequences. J Cell Sci, 2017. 130(2): p. 372-381. 74. Kida, Y., et al., Stability and flexibility of marginally hydrophobic- segment stalling at the endoplasmic reticulum translocon. Mol Biol Cell, 2016. 27(6): p. 930-40. 75. von Heijne, G., Control of topology and mode of assembly of a polytopic membrane protein by positively charged residues. Nature, 1989. 341(6241): p. 456-8.

76. Braakman, I. and D.N. Hebert, Protein folding in the endoplasmic reticulum. Cold Spring Harb Perspect Biol, 2013. 5(5): p. a013201. 77. Lamriben, L., et al., N-Glycan-based ER Molecular Chaperone and Protein Quality Control System: The Calnexin Binding Cycle. Traffic, 2016. 17(4): p. 308-26. 78. Nilsson, I.M. and G. von Heijne, Determination of the distance between the oligosaccharyltransferase active site and the endoplasmic reticulum membrane. J Biol Chem, 1993. 268(8): p. 5798-801. 79. Kaplan, H.A., J.K. Welply, and W.J. Lennarz, Oligosaccharyl transferase: the central enzyme in the pathway of glycoprotein assembly. Biochim Biophys Acta, 1987. 906(2): p. 161-73. 80. Wallis, A.K. and R.B. Freedman, Assisting oxidative protein folding: how do protein disulphide-isomerases couple conformational and chemical processes in protein folding? Top Curr Chem, 2013. 328: p. 1-34. 81. Chen, W., et al., Cotranslational folding and calnexin binding during glycoprotein synthesis. Proc Natl Acad Sci U S A, 1995. 92(14): p. 6229-33. 82. Wang, N., et al., The cotranslational maturation program for the type II membrane glycoprotein influenza neuraminidase. J Biol Chem, 2008. 283(49): p. 33826-37. 83. Daniels, R., et al., N-linked glycans direct the cotranslational folding pathway of influenza hemagglutinin. Mol Cell, 2003. 11(1): p. 79-90. 84. Hogue, B.G. and D.P. Nayak, Synthesis and processing of the influenza virus neuraminidase, a type II transmembrane glycoprotein. Virology, 1992. 188(2): p. 510-7. 85. Hebert, D.N., B. Foellmer, and A. Helenius, Calnexin and calreticulin promote folding, delay oligomerization and suppress degradation of influenza hemagglutinin in microsomes. EMBO J, 1996. 15(12): p. 2961-8. 86. Tatu, U., C. Hammond, and A. Helenius, Folding and oligomerization of influenza hemagglutinin in the ER and the intermediate compartment. EMBO J, 1995. 14(7): p. 1340-8. 87. Saito, T., G. Taylor, and R.G. Webster, Steps in maturation of influenza A virus neuraminidase. J Virol, 1995. 69(8): p. 5011-7. 88. da Silva, D.V., et al., Assembly of subtype 1 influenza neuraminidase is driven by both the transmembrane and head domains. J Biol Chem, 2013. 288(1): p. 644-53. 89. Nordholm, J., et al., Polar residues and their positional context dictate the transmembrane domain interactions of influenza A neuraminidases. J Biol Chem, 2013. 288(15): p. 10652-60. 90. Air, G.M., Influenza neuraminidase. Influenza Other Respir Viruses, 2012. 6(4): p. 245-56.

91. Chong, A.K., M.S. Pegg, and M. von Itzstein, Influenza virus sialidase: effect of calcium on steady-state kinetic parameters. Biochim Biophys Acta, 1991. 1077(1): p. 65-71. 92. Colacino, J.M., et al., A single sequence change destabilizes the influenza virus neuraminidase tetramer. Virology, 1997. 236(1): p. 66-75. 93. Vavricka, C.J., et al., Structural and functional analysis of laninamivir and its octanoate prodrug reveals group specific mechanisms for influenza NA inhibition. PLoS Pathog, 2011. 7(10): p. e1002249. 94. Shtyrya, Y.A., L.V. Mochalova, and N.V. Bovin, Influenza virus neuraminidase: structure and function. Acta Naturae, 2009. 1(2): p. 26-32. 95. Cohen, M., et al., Influenza A penetrates host mucus by cleaving sialic acids with neuraminidase. Virol J, 2013. 10: p. 321. 96. Matrosovich, M.N., et al., Neuraminidase is important for the initiation of influenza virus infection in human airway epithelium. J Virol, 2004. 78(22): p. 12665-7. 97. Matrosovich, M.N., et al., Human and avian influenza viruses target different cell types in cultures of human airway epithelium. Proc Natl Acad Sci U S A, 2004. 101(13): p. 4620-4. 98. Webster, R.G. and W.G. Laver, Preparation and properties of antibody directed specifically against the neuraminidase of influenza virus. J Immunol, 1967. 99(1): p. 49-55. 99. Palese, P. and J. Schulman, Isolation and characterization of influenza virus recombinants with high and low neuraminidase activity. Use of 2-(3'-methoxyphenyl)-n-acetylneuraminic acid to identify cloned populations. Virology, 1974. 57(1): p. 227-37. 100. Palese, P., et al., Characterization of temperature sensitive influenza virus mutants defective in neuraminidase. Virology, 1974. 61(2): p. 397-410. 101. Lakdawala, S.S., et al., Eurasian-origin gene segments contribute to the transmissibility, aerosol release, and morphology of the 2009 pandemic H1N1 influenza virus. PLoS Pathog, 2011. 7(12): p. e1002443. 102. Zanin, M., et al., Pandemic Swine H1N1 Influenza Viruses with Almost Undetectable Neuraminidase Activity Are Not Transmitted via Aerosols in Ferrets and Are Inhibited by Human Mucus but Not Swine Mucus. J Virol, 2015. 89(11): p. 5935-48. 103. von Itzstein, M., et al., Rational design of potent sialidase-based inhibitors of influenza virus replication. Nature, 1993. 363(6428): p. 418-23. 104. Hanessian, S., et al., Design, synthesis, and neuraminidase inhibitory activity of GS-4071 analogues that utilize a novel hydrophobic paradigm. Bioorg Med Chem Lett, 2002. 12(23): p. 3425-9.

105. Chen, Y.Q., et al., Influenza Infection in Humans Induces Broadly Cross-Reactive and Protective Neuraminidase-Reactive Antibodies. Cell, 2018. 173(2): p. 417-429 e10. 106. Dowdle, W.R., et al., Inactivated influenza vaccines. 2. Laboratory indices of protection. Postgrad Med J, 1973. 49(569): p. 159-63. 107. Ogra, P.L., et al., Clinical and immunologic evaluation of neuraminidase-specific influenza A virus vaccine in humans. J Infect Dis, 1977. 135(4): p. 499-506. 108. Monto, A.S., et al., Antibody to Influenza Virus Neuraminidase: An Independent Correlate of Protection. J Infect Dis, 2015. 212(8): p. 1191-9. 109. Couch, R.B., et al., Antibody correlates and predictors of immunity to naturally occurring influenza in humans and the importance of antibody to the neuraminidase. J Infect Dis, 2013. 207(6): p. 974-81. 110. Schulman, J.L., M. Khakpour, and E.D. Kilbourne, Protective effects of specific immunity to viral neuraminidase on influenza virus infection of mice. J Virol, 1968. 2(8): p. 778-86. 111. Wohlbold, T.J., et al., Broadly protective murine monoclonal antibodies against influenza B virus target highly conserved neuraminidase epitopes. Nat Microbiol, 2017. 2(10): p. 1415-1424. 112. Wan, H., et al., Structural characterization of a protective epitope spanning A(H1N1)pdm09 influenza virus neuraminidase monomers. Nat Commun, 2015. 6: p. 6114. 113. Wan, H., et al., Molecular basis for broad neuraminidase immunity: conserved epitopes in seasonal and pandemic H1N1 as well as H5N1 influenza viruses. J Virol, 2013. 87(16): p. 9290-300. 114. Marcelin, G., et al., A contributing role for anti-neuraminidase antibodies on immunity to pandemic H1N1 2009 influenza A virus. PLoS One, 2011. 6(10): p. e26335. 115. Walz, L., et al., Neuraminidase-Inhibiting Antibody Titers Correlate with Protection from Heterologous Influenza Virus Strains of the Same Neuraminidase Subtype. J Virol, 2018. 92(17). 116. Rockman, S., et al., Neuraminidase-inhibiting antibody is a correlate of cross-protection against lethal H5N1 influenza virus in ferrets immunized with seasonal influenza vaccine. J Virol, 2013. 87(6): p. 3053-61.