Structures_and_Functions_of_Retroviral_RNAs_The_Multiple_Facets.pdf

Structures and Functions of Retroviral RNAs

Nucleic Acids Set
coordinated by
Marie-Christine Maurel
Volume 1
Structures and Functions
of Retroviral RNAs
The Multiple Facets of the
Retroviral Genome
Philippe Fossé

First published 2022 in Great Britain and the United States by ISTE Ltd and John Wiley & Sons, Inc.
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Contents
Foreword . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . vii
Marylène MOUGEL
Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xi
Chapter 1. General Information on Retroviruses . . . . . . . . . . . . . 1
1.1. Common characteristics of retroviruses . . . . . . . . . . . . . . . . . . . 1
1.1.1. Untranslated regions . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2
1.1.2. Translated regions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3
1.2. Architecture of the virion. . . . . . . . . . . . . . . . . . . . . . . . . . . . 5
1.3. Replication cycle of retroviruses . . . . . . . . . . . . . . . . . . . . . . . 6
1.3.1. Early phase. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7
1.3.2. Late phase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9
Chapter 2. Effects of the Structure of Retroviral RNA on Reverse
Transcription . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11
2.1. Reverse transcription of genomic RNA . . . . . . . . . . . . . . . . . . . 11
2.2. RNA structures involved in the initiation of reverse transcription. . . . 14
2.2.1. A cellular tRNA as RT primer . . . . . . . . . . . . . . . . . . . . . . 14
2.2.2. Formation of the template–primer duplex . . . . . . . . . . . . . . . 16
2.2.3. Role of the structure of the template–primer duplex
upon initiation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19
2.3. RNA structures involved in the first strand transfer . . . . . . . . . . . . 21
2.3.1. Actors involved in the first strand transfer . . . . . . . . . . . . . . . 21
2.3.2. Molecular basis of R–r pairing . . . . . . . . . . . . . . . . . . . . . . 22
2.4. RNA structures promoting genetic recombination. . . . . . . . . . . . 26
2.4.1. Internal strand transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . 26
2.4.2. RNA structures triggering internal strand transfer. . . . . . . . . . . 28

vi Structures and Functions of Retroviral RNAs
Chapter 3. RNA Structures Regulating the Expression of the
Retroviral Genome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31
3.1. Regulatory RNA structures of proviral DNA transcription. . . . . . . . 31
3.1.1. The transcriptional activator Tat . . . . . . . . . . . . . . . . . . . . . 31
3.1.2. TAR structures with one stem-loop . . . . . . . . . . . . . . . . . . . 33
3.1.3. TAR structures with two stem-loops . . . . . . . . . . . . . . . . . . 37
3.2. RNA structures regulating genomic RNA maturation . . . . . . . . . . . 38
3.2.1. The negative regulator of splicing of RSV . . . . . . . . . . . . . . . 39
3.2.2. Structural diversity and alternative splicing in HIV-1 . . . . . . . . 41
3.3. RNA structures regulating the export of retroviral RNAs. . . . . . . . . 44
3.3.1. Export of unspliced RNA in simple retroviruses . . . . . . . . . . . 44
3.3.2. Unspliced RNA export in complex retroviruses . . . . . . . . . . . . 53
3.4. RNA structures regulating the translation of retroviral RNAs . . . . . . 60
3.4.1. IRESs of simple retroviruses . . . . . . . . . . . . . . . . . . . . . . . 61
3.4.2. Translation initiation in complex retroviruses . . . . . . . . . . . . . 63
Chapter 4. Encapsidation of Genomic RNA in the
Retroviral Particle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73
4.1. RNA structures and mechanisms governing gRNA dimerization . . . . 73
4.1.1. Structures and mechanisms in alpharetroviruses . . . . . . . . . . . 74
4.1.2. Structures and mechanisms in betaretroviruses . . . . . . . . . . . . 77
4.1.3. Structures and mechanisms in deltaretroviruses . . . . . . . . . . . . 79
4.1.4. Structures and mechanisms in gammaretroviruses . . . . . . . . . . 81
4.1.5. Structures and mechanisms in lentiviruses . . . . . . . . . . . . . . . 85
4.2. RNA structures and mechanisms regulating gRNA encapsidation . . . 96
4.2.1. Structures and mechanisms in alpharetroviruses . . . . . . . . . . . 96
4.2.2. Structures and mechanisms in betaretroviruses . . . . . . . . . . . . 99
4.2.3. Structures and mechanisms in deltaretroviruses . . . . . . . . . . . . 104
4.2.4. Structures and mechanisms in gammaretroviruses . . . . . . . . . . 106
4.2.5. Structures and mechanisms in lentiviruses . . . . . . . . . . . . . . . 108
Conclusion. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115
Glossary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117
List of Acronyms. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 121
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 127
Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 159

Foreword
The sequencing of the human genome has turned our vision of biology
upside down, revealing little by little the complexity of living organisms. We
have realized that genes, DNA coding for proteins, do not control everything
in the cell. Of the two meters of DNA in a cell, the genes represent only six
centimeters. However, almost all of this DNA (80–90%) is transcribed into
RNA. When they are not coding, these RNAs are regulators of the cell, and
their role is strongly dependent on their structures. Unlike DNA, RNA is a
dynamic molecule that can adopt different conformations, giving it several
functions.
Sequencing has also revealed the strong presence of sequences of
retroviral origin in the form of retroelements, retrotransposons or
endogenous retroviruses, which are vestiges of our cohabitation with these
viruses. Their roles remain mostly unknown: some are beneficial, for
example by helping the placenta to form, and others trigger pathological
processes, in particular cancerous ones. When these are able to leave the cell
and infect other cells, they are called retroviruses. With the arrival of the
Covid-19 pandemic caused by SARS-CoV-2, RNA viruses are now
perceived as a real threat on a global scale. This pandemic is reminiscent of
other pandemics, including acquired immunodeficiency syndrome (AIDS)
caused by the human immunodeficiency virus (HIV) retrovirus. These
pandemics highlight the strong need to understand the functioning
mechanisms of these viruses and their ability to spread.

viii Structures and Functions of Retroviral RNAs
RNA, the probable ancestor of DNA and proteins, is a fascinating
molecule because of its complexity, multifunctionality and remarkable
adaptation to its host. This versatility of RNA is consistent with the multiple
primary, secondary and tertiary structures it can adopt. Retroviral RNA is a
perfect example with its numerous functions. Indeed, it plays several roles
during infection: initially, it serves as a template for DNA synthesis through
the reverse transcriptase, then as messenger RNA (mRNA) encoding
structural and enzymatic proteins, and finally as the genome, the guardian of
the virus genetic information (gRNA). It is able to hijack the cellular
machinery of its host and transgress the rules of cellular RNA metabolism.
For example, when used as mRNA, it can escape splicing and leave the
nucleus despite the persistence of introns. It can also take two different
export routes to exit the nucleus, presumably in different conformations.
Recent studies have led to the discovery and understanding of the
mechanisms of new nuclear export pathways, which are also used by some
cellular RNAs. Retroviral RNAs have assisted in major discoveries and have
shattered several biological dogmas.
This book reveals how RNA folds back on itself when it comes into
contact with its partners encountered during the different stages of virus
replication in the cell. We will learn how the structure of RNA evolves when
it is copied into DNA by the viral reverse transcriptase after the virus enters
the cell. One of the major questions of retrovirology is understanding how all
the viral components (proteins and RNA), present in multiple copies, are
found in a concerted manner, in number and time, at the periphery of the
cell, in order to assemble and form new viruses, ready to disseminate outside
the cell. To be infectious, a virus must contain not one but two copies of the
same gRNA; we then speak of dimeric gRNA. How, among the multitude of
cellular RNAs present in the cytoplasm of the cell, is the gRNA able to pair
up and find its protein partners to form new viruses? Once again, the answer
lies in the structure of this RNA.
Philippe Fossé, a distinguished director of research at the CNRS, focuses
all his research on the study of the RNA structure of various retroviruses
(avian, murine and human), with the aim of understanding the complex
relationships between RNA structure and function. This is very meticulous
work on a molecular scale, which requires extreme rigor and a certain

Foreword ix
insight acquired through experience. Indeed, computer programs for
structure prediction, although increasingly sophisticated, are not by
themselves sufficient to account for the versatility and folding dynamics of
these RNAs.
Marylène MOUGEL
Institut de recherche en infectiologie
de Montpellier (IRIM)
May 2022

Preface
Retroviruses, which are single-stranded RNA viruses of positive polarity,
have been identified in various vertebrate groups but not in invertebrates.
Retroviruses originated with their aquatic vertebrate hosts at least 450
million years ago and have evolved through interactions with them.
Retroviruses have contributed to vertebrate evolutionary processes. The most
prominent example in host evolution is the formation of the placenta in the
ancestors of placental mammals through several independent retroviral
infections. Retroviruses are divided into two subfamilies (Orthoretrovirinae
and Spumaretrovirinae). Spumaretroviruses infect a wide variety of
mammals and are generally non-pathogenic. In contrast, retroviruses
belonging to the Orthoretrovirinae subfamily are often responsible for
pathologies in the vertebrates they infect. From the beginning of the 20th
century until the beginning of the 1980s, in order to elucidate the
mechanisms of carcinogenesis, numerous studies were focused on avian and
murine retroviruses that induce leukemia and cancer in their hosts. These
studies led to the discovery of oncogenes, contributing to the understanding
of the regulation of eukaryotic gene expression and the characterization of
the stages of the retrovirus replicative cycle. In addition, they have enabled
fundamental scientific and technological advances in biology through the
discovery of reverse transcriptase and the use of retroviral vectors in the
analysis of gene expression. Since its discovery in 1983, HIV, the causative
agent of AIDS, has been the main focus of retrovirology research.

xii Structures and Functions of Retroviral RNAs
Although the retrovirus genome is small (7–12 kb), in its RNA form, it
performs multiple functions other than serving as a messenger for the
synthesis of proteins necessary for the production of infectious viral
particles. These functions, some of which vary according to the retroviral
species, depend mainly on the structures adopted by retroviral RNA. In this
book, which is based on the extensive scientific literature, I provide a
non-exhaustive review of the knowledge acquired on the structure–function
relationships of RNA in different retroviral species. In Chapter 1, I present
general knowledge on retroviruses, as it is necessary to understand the
molecular mechanisms regulated by RNA structures. Reverse transcription
of retroviral RNA is a complex and essential process in the replicative cycle
of retroviruses. The key steps of this process, which rely on interactions
involving DNA and RNA structures, are presented in Chapter 2. Chapter 3
shows that, in some retroviruses, their genomic RNA forms secondary
structures that serve as signals to regulate proviral DNA transcription,
maturation, export and translation of retroviral RNA. Encapsidation is a
process common to all retroviruses that allows the incorporation of two
genomic RNA molecules into the viral particle. It requires interactions
between several molecules of a retroviral protein and secondary structures of
the genomic RNA, as described in Chapter 4.
For ease of reading, a glossary and a list of abbreviations and acronyms
are included following the conclusion of this book. I hope that this book
enables students and researchers to understand the multiple facets of
retroviral RNA and contributes to developing their knowledge and critical
thinking in the fields of research involving functional RNAs.
May 2022

1
General Information on Retroviruses
1.1. Common characteristics of retroviruses
Retroviruses, like all viruses, are parasites that lack the genetic
information encoding the enzymes of intermediary metabolism and can
therefore only replicate inside living cells. They infect vertebrates and can
cause cancerous tumors, leukemia, neurological disorders and AIDS. Most
retroviruses are exogenous and their transmission is achieved by contagion
between distinct individuals. Others, known as endogenous, are integrated
into the host genome and are transmitted hereditarily.
Although they are capable of infecting different animal host cells and
causing different pathologies, all retroviruses have common structural and
functional characteristics that allow them to be grouped in the family
Retroviridae (see Table 1.1). In all retroviruses, the genetic information is
carried by a single-stranded RNA.
The term retrovirus comes from the fact that their replication cycle
imposes a passage from the RNA genome to a DNA form; retro- thus refers
to the unusual direction from RNA to DNA. This passage is carried out by
way of reverse transcriptase (RT), a retroviral enzyme that is an RNA- and
DNA-dependent DNA polymerase (Baltimore 1970; Temin and Mizutani
1970).
Structures and Functions of Retroviral RNAs: The Multiple Facets of the Retroviral Genome,
First Edition. Philippe Fossé.
© ISTE Ltd 2022. Published by ISTE Ltd and John Wiley & Sons, Inc.

2 Structures and Functions of Retroviral RNAs
The RNA genome of retroviruses, called genomic RNA (gRNA), consists
of terminal non-coding regions necessary for viral replication and internal
regions that code for viral enzymes and structural proteins (see Figure 1.1).
In the following, the generic term retrovirus only refers to retroviruses
belonging to the subfamily Orthoretrovirinae.
Family Subfamily Type Species
Retroviridae Orthoretrovirinae
Alpharetrovirus
Betaretrovirus
Deltaretrovirus
Espsilonretrovirus
Gammetrovirus
Lentivirus
9 species
5 species
4 species
3 species
18 species
10 species
Spumaretrovirinae Spumavirus 19 species
Table 1.1. Classification of retroviruses
1
1.1.1. Untranslated regions
The untranslated terminal regions of gRNA are arranged in the same
order in all retroviruses (see Figure 1.1), but depending on the species, they
differ in terms of size, structure and some of their characteristics (Coffin
1992).
gRNA is derived from the same maturation process as cellular RNAs. A
methylated guanosine m7G5' ppp5' is present at the 5' end of the 5'
non-coding region; this cap is required for ribosome attachment and is
important for the translation of viral RNAs that serve as messengers
(Bolinger and Boris-Lawrie 2009). A poly(A) tail consisting of
approximately 200 adenines is present at the 3' end of the 3' non-coding
region (Vogt 1997).
1.1.1.1. The 5' untranslated region (5'-UTR)
This region is composed of four sequences (R, U5, PBS and L). The R
(repeat) sequence, 16–247 nucleotides depending on the retrovirus, is present
1 Available from: https://talk.ictvonline.org/ictv-reports/ictv_9th_report/reverse-transcribing-
dna-and-rna-viruses-2011/w/rt_viruses/161/retroviridae.

General Information on Retroviruses 3
at both ends in all retroviruses (Klaver and Berkhout 1994). It plays an
essential role in the replicative strategy of the retrovirus. The unique U5
sequence, 80–240 nucleotides depending on the retrovirus, is located
downstream of the R sequence (Vogt 1997). It is the first region of the
gRNA that is copied into DNA during reverse transcription. The primer
binding site (PBS) sequence, consisting of 18 nucleotides, pairs with the 3'
end of a cellular tRNA molecule that serves as a primer to start reverse
transcription (Vogt 1997). The nature of the tRNA primer differs among
retroviruses. The leader (L) sequence lies between the PBS and the gag gene
and comprises at least part of the gRNA packaging signal in the virus
particle.
Figure 1.1. Genetic organization of retroviruses. (a) Genomic RNA and
(b) proviral DNA. The cellular DNA is represented by a wavy line. For a
color version of this figure, see www.iste.co.uk/fosse/structures.zip
1.1.1.2. The 3' untranslated region (3'-UTR)
This region is composed of three sequences (PPT, U3 and R). The
polypurine tract (PPT) sequence, very rich in purines, 9–15 depending on the
retrovirus, is located upstream of the U3 sequence. It serves as a primer for
the synthesis of the (+) strand of the proviral DNA (Vogt 1997). The U3
sequence, a unique 3' sequence, is located between the PPT and the R
sequence. U3 contains the promoter and regulator elements of viral RNA
transcription. The R sequence is identical to that present in the 5'-UTR. It
enables the first strand transfer, which is a crucial step in reverse
transcription.
1.1.2. Translated regions
All retroviruses have the gag, pol and env genes (see Figure 1.1). The gag
cistron and the pol cistron are often considered a single cistron, although
there may be a phase shift between the two reading frames.
a)
b)

1.1.2.1. The gag gene (group-specific antigen)
This codes for a polyprotein precursor. This precursor is translated from
gRNA which is an unspliced mRNA. During the maturation of the virus
particle, the Gag precursor is cleaved by the viral protease (PR) to generate
the structural proteins MA, CA and NC (see Figure 1.2).
1.1.2.2. The pol (polymerase) gene
The Gag–Pol polypeptide precursor is translated from gRNA when
protein synthesis is not stopped at the stop codon of the gag gene.
Depending on the retrovirus, two types of mechanisms can be used to
continue translation after the stop codon: a reading frame-shift mechanism
between the gag and pol genes (Jacks and Varmus 1985; Moore et al. 1987;
Wilson et al. 1988; Nam et al. 1993) or the incorporation of an amino acid at
the stop codon (Yoshinaka et al. 1985a; Yoshinaka et al. 1985b). The rate of
synthesis of the Gag–Pol precursor is about 5% compared to that of the Gag
precursor (Shehu-Xhilaga et al. 2001a). Proteolysis of Gag–Pol by PR is the
origin of the structural proteins (MA, CA and NC) and viral enzymes: RT
and integrase (IN) (Konvalinka et al. 2015).
Figure 1.2. Products of the gag, pol and env genes. For a color
version of this figure, see www.iste.co.uk/fosse/structures.zip
COMMENTARY ON FIGURE 1.2.– Schematic representation of the three
polypeptide precursors synthesized from the gag, pol and env genes.
Although in all retroviruses the protease-encoding sequence (PR) is always
between the gag and pol genes, its reading frame varies among retroviral
Gag-Pol MA CA NC
gag
PR RT IN
MA CA NC
Gag
pol
Env SU TM
env

species (Konvalinka et al. 2015). For example, in HIV-1, PR is in the same
reading frame as the pol gene, whereas it is in the reading frame of gag in
alpharetroviruses and a separate reading frame in Mason–Pfizer monkey
virus (MPMV).
1.1.2.3. The env (envelope) gene
The env gene, unlike the gag and pol genes, is translated from spliced
mRNA. The Env polypeptide precursor undergoes post-translational
modifications, such as glycosylation, before being cleaved by a cellular
protease into two subunits: the surface protein (SU) and the transmembrane
protein (TM). The two subunits remain associated by non-covalent bonds in
most retroviruses and form a trimer, with the portion exposed on the surface
of the virus particle constituting the spike involved in host cell recognition
(Steckbeck et al. 2014).
1.2. Architecture of the virion
In retroviruses, the infectious viral particle called the virion is spherical,
100–200 nm in diameter (Zhang et al. 2015). Retroviruses are enveloped
viruses because the outer envelope of the virion is composed of a lipid
bilayer. This is derived from the plasma membrane of the infected cell and is
enriched in viral envelope protein (see Figure 1.3). The inner part of the
virus forms a shell called the capsid, which is made up of self-assembling
capsid proteins (CA). In all retroviruses belonging to the Orthoretrovirinae
subfamily, the capsid contains the viral enzymes (PR, RT and IN) and the
diploid genome of the virus consisting of two gRNA molecules in close
association with the nucleocapsid (NC) proteins (Vogt 1997). The capsid
also contains mRNA, tRNA and host cell proteins. The matrix consists
mainly of matrix proteins (MA) and is located between the envelope and the
capsid.

Figure 1.3. Schematic representation of the virion. The structural organization of the
virion is the same in all Orthoretrovirinae. The capsid containing viral enzymes and
gRNA associated with NC molecules forms a closed shell with a morphology that
varies among retroviral species (conical, tubular, nearly spherical or polyhedral)
(Zhang et al. 2015). For a color version of this figure, see www.iste.co.uk/fosse/
structures.zip
1.3. Replication cycle of retroviruses
All retroviruses have an infection cycle with common stages. The
replicative cycle of a retrovirus is divided into two main phases (D’Souza
and Summers 2005): early and late (see Figure 1.4). The early phase

includes the steps from binding the virion to the host cell receptor up to the
integration of the double-stranded viral DNA into the host cell genome
(proviral DNA). The late phase includes the following steps from
transcription of the proviral DNA to the release of new virions into the
extracellular medium.
1.3.1. Early phase
1.3.1.1. Entry of the virus into the host cell
The attachment of the virion to the target cell is mediated by a specific
interaction between a receptor on the target cell and the envelope
glycoprotein of the virus. There are several types of cellular receptors
recognized by retroviruses; however, only one type of receptor is used by a
retroviral species (Sommerfelt 1999). The envelope glycoprotein–receptor
interaction triggers fusion of the cell and viral membranes and thus leads to
the release of the capsid into the cytoplasm of the host cell (Lindemann et al.
2013). In the case of retroviruses belonging to the genus Lentivirus, such as
HIV type 1 (HIV-1) and type 2 (HIV-2), an interaction of the envelope
glycoprotein with a coreceptor is required for strong binding of the virion to
the cell membrane and to allow fusion of the cell and viral membranes
(Deng et al. 1996; Feng et al. 1996).
1.3.1.2. Reverse transcription and decapsidation
The reverse transcription of the gRNA, which is described in detail in
Chapter 2, begins as soon as the capsid enters the cell. This complex process
converts the single-stranded gRNA into double-stranded DNA, with a long
repeating sequence (LTR) at both ends (see Figure 1.1). Capsid disassembly
(decapsidation) has primarily been studied in HIV-1 and is partly dependent
on reverse transcription (Dismuke and Aiken 2006; Arhel et al. 2007; Hulme
et al. 2011; Cosnefroy et al. 2016; Mamede et al. 2017; Fernandez et al.
2019). There are three models of decapsidation:
– progressive decapsidation in the cytoplasm during transport to the
nuclear membrane (Mamede et al. 2017);
– decapsidation at a nuclear pore (Fernandez et al. 2019);
– decapsidation after entering the nucleus of an intact or nearly intact
capsid (Burdick et al. 2020).

Figure 1.4. Replicative cycle of a retrovirus. For a color version
of this figure, see www.iste.co.uk/fosse/structures.zip
COMMENTARY ON FIGURE 1.4. – (1) Attachment of the virion to the target
cell. (2) Fusion of cell and viral membranes. (3) Reverse transcription and
transport of the capsid to the nuclear membrane. (4) Decapsidation at a
nuclear pore. (5) Entry of double-stranded viral DNA (orange band) into the
nucleus and integration into cellular DNA (gray band). (6) Transcription of
the proviral DNA. (7) Splicing of a fraction of the gRNA molecules.
(8) Nuclear export of a fraction of the gRNA molecules. (9) Nuclear export
of the viral mRNA. (10) Translation of viral mRNAs. (11) Translation of
gRNA molecules. (12) Assembly of a viral particle involving two gRNA
molecules, SU and TM proteins and Gag and Gag–Pol precursors.
(13) Budding of the viral particle. (14) Release of an immature virus particle
into the extracellular environment. (15) Maturation of the viral particle.

1.3.1.3. Integration of the viral genome into the DNA of the host cell
The integration of double-stranded viral DNA into the host cell genome,
leading to proviral DNA (see Figure 1.1), is an essential step in the
replicative cycle of retroviruses. It allows the virus, on the one hand, to
express its genes using the cellular machinery of transcription and splicing,
and on the other hand, to maintain its genome in the cells during cell
divisions. This integration process requires IN, an enzyme encoded by the
retrovirus pol gene, and cellular enzymes (Ciuffi 2016).
1.3.2. Late phase
1.3.2.1. Synthesis, maturation and export of viral RNA
Once integrated into the cellular genome, the proviral DNA is transcribed
into RNA by the cell’s transcription machinery. An m7G5'ppp5' cap and a
poly(A) tail are added to the 5' and 3' ends of the newly synthesized RNA,
respectively, corresponding to the retrovirus gRNA (Vogt 1997; Bolinger
and Boris-Lawrie 2009). A fraction of the gRNA population is not spliced
and is exported to the cytoplasm. From a fraction of the gRNA population,
the cellular splicing machinery generates viral mRNAs that are exported to
the cytoplasm. In all retroviruses, the gRNA contains at least one intron
bounded by a splice donor site and a splice acceptor site to generate the env
mRNA. The number of mRNA isoforms generated by alternative splicing of
the gRNA varies among retroviruses. In HIV-1, alternative splicing can
generate up to 100 multi-spliced RNAs (Ocwieja et al. 2012). The role of
RNA structures in viral RNA synthesis, maturation and export is discussed
in detail in Chapter 3.
1.3.2.2. Translation of viral RNA
After being exported to the cytoplasm, a fraction of the gRNA population
is encapsidated into a forming virus particle and/or translated into Gag and
Gag–Pol polypeptide precursors, which participate in virion formation. The
SU and TM subunits of the envelope glycoprotein are synthesized from the
env mRNA and integrated into the virus particle. Some of the proteins
synthesized from the other viral mRNAs are also incorporated into the virion
(e.g. Vpr in the case of lentiviruses) (Fabryova and Strebel 2019). The RNA
structures and mechanisms that regulate protein synthesis are described in
Chapter 3.

1.3.2.3. Formation of the virion
Two gRNA molecules are specifically encapsidated in the budding virus
particle (see Figure 1.4). The encapsidation of the viral RNA, which requires
the Gag precursor, is an obligatory step in the infectious cycle (D’Souza and
Summers 2005). The RNA structures and mechanisms involved in this step
are described in Chapter 4. The gRNA and Gag polypeptide play an
important role in the assembly of new virus particles (D’Souza and Summers
2005; Olson et al. 2015). With the exception of betaretroviruses, assembly
and budding of a new viral particle occur simultaneously at the plasma
membrane (D’Souza and Summers 2005; Prchal et al. 2013). Gag and
Gag–Pol polypeptides bind to the plasma membrane through their
N-terminus. The presence at the plasma membrane of the Gag, Gag–Pol
precursors, SU and TM proteins and two copies of the gRNA triggers the
assembly and budding of a new virus particle. The virus particle is not
immediately infectious after being released into the extracellular medium.
It undergoes a maturation process that allows it to become infectious
(virion). The maturation process is initiated by the viral PR which cleaves
the Gag and Gag–Pol precursors. It begins during or just after the release of
the virus particle (Pornillos and Ganser-Pornillos 2019).

2
Effects of the Structure of Retroviral
RNA on Reverse Transcription
2.1. Reverse transcription of genomic RNA
Thanks to its DNA/RNA-dependent DNA polymerase activity, RT is the
main actor in the reverse transcription of genomic RNA. For this reason, it is
one of the main targets of pharmacological agents used against HIV-1. RT
has no proofreading activity. Therefore, there is a particularly high rate of
mutation during reverse transcription. In conjunction with the recombination
events occurring during reverse transcription, mutations give retroviruses a
high degree of genetic variability that leads them to evade host immune
defenses and develop resistance to antiretroviral agents. The NC is the other
viral protein that plays an important role in reverse transcription (Levin et al.
2010). This protein, which is not an enzyme, is rich in basic amino acids,
and has one or two zinc fingers depending on the retroviral species (see
Figure 2.1). It is a chaperone protein of nucleic acids, that is, it allows a
nucleic acid to form the secondary structure that is thermodynamically most
stable, which generally corresponds to the conformation with the maximum
number of base pairs (Thomas and Gorelick 2008; Darlix et al. 2011). The
chaperone activity of the NC is the result of three properties (Levin et al.
2010):
– nucleic acid aggregation, which is mainly due to basic amino acids;
– destabilization of short nucleic acid duplexes by zinc fingers;
– short residence time on the nucleic acid.

Figure 2.1. Examples of NC sequences. NCs are small proteins (less than 100
amino acids) that have one (for example MoMuLV) or two (for example RSV and
HIV-1) zinc fingers (source: Genbank: MoMuLV (J02255), HIV-1 (AF324493), RSV
(J02342))
Although in a few virions, reverse transcription of gRNA is initiated
before entry into the cell, and it primarily begins when the infectious particle
enters the cell (Trono 1992; Zhang et al. 1993; Hooker and Harrich 2003).
The reverse transcription of the single-stranded gRNA is characterized by a
sequence of steps leading to the synthesis of double-stranded DNA with an
LTR at each end (Gilboa et al. 1979) (see Figure 2.2). Reverse transcription
of a single gRNA molecule is theoretically sufficient to produce a double-
stranded DNA molecule. However, studies suggest that the two gRNA
molecules present as a dimer in the capsid are used to generate a single
molecule of a double-stranded DNA molecule (Panganiban and Fiore 1988;
Yu et al. 1998). The RNA structures that are involved in certain steps of
reverse transcription will be described below.

Effects of the Structure of Retroviral RNA on Reverse Transcription 13
Figure 2.2. Reverse transcription of the retroviral genome. For a color
COMMENTARY ON FIGURE 2.2 – In this diagram, only one of the two strands
of the gRNA dimer is shown to facilitate the understanding of the different
steps. (1) RT synthesis of negative polarity ssDNA from the tRNA primer
bound to the PBS sequence. (2) RT RNase H degradation of the copied R-U5

region and first strand transfer at the R–r sequences. (3) Synthesis of the
negative polarity DNA strand and degradation of the copied RNA strand by
RT RNase H activity. (4) Further synthesis of the negative polarity DNA
strand and initiation of the positive polarity DNA strand synthesis from the
PPT RNA primer. (5) Complete synthesis of the negative polarity DNA. (6)
Second strand transfer at the PBS–pbs complementary sequences. (7) The
(+) and (-) DNA strands are extended by RT to produce double-stranded
retroviral DNA with an LTR at each end. The retroviral genomic RNA is
shown as a blue line. The tRNA primer is shown with a purple line. The DNA
strands (-) and (+) are, respectively, shown with brown and orange lines.
The direction of RT polymerization is shown with an arrowhead. The
degradation of RNA by the RNase H activity of RT is shown with a dashed
line. Sequence names are written in capital letters for the (+) strand and in
lower case for the (-) strand.
2.2. RNA structures involved in the initiation of reverse
transcription
2.2.1. A cellular tRNA as RT primer
Like all DNA polymerases, RT needs a primer to start DNA
polymerization. In retroviruses, the primer is a cellular tRNA (Marquet et al.
1995) which is tRNALys3
in lentiviruses (e.g. HIV-1), tRNATrp
in
alpharetroviruses (e.g. Rous sarcoma virus (RSV)) and tRNAPro
in murine
retroviruses (e.g. Moloney murine leukemia virus (MoMuLV)) (see
Figure 2.3).
The tRNA primer is incorporated into the virion as it is assembled.
Incorporation of tRNA can be highly selective (e.g. in RSV and HIV-1) or
minimally selective (e.g. in MoMuLV) (Mak and Kleiman 1997). Selective
incorporation has been extensively studied in HIV-1; this involves the
polypeptide precursors Gag and Gag–Pol, and Lysyl–tRNA synthetases of
the host cell (Kaminska et al. 2007; Kleiman et al. 2010).

Figure 2.3. Examples of secondary structures of tRNAs and gRNAs
involved in the PBS–primer interaction. For a color version
COMMENTARY ON FIGURE 2.3.– For each of the three retroviruses taken as
examples, the secondary structure of the tRNA primer it uses is shown in the
upper part of the figure, while the lower part shows the secondary structure
of the portion of the gRNA that contains the PBS sequence (Keith and
Heyman 1990; Hackett et al. 1991; Baudin et al. 1993; Mougel et al. 1993).
Complementary sequences consisting of 18 nucleotides (PBS in viral RNA

and anti-PBS in tRNA primer) are purple and blue, respectively. Letters not
corresponding to A, C, G and U correspond to modified nucleotides that are
frequent in tRNAs. The arms (acceptor, T, anticodon and D) are indicated by
the letters in orange (Ac, T, A and D, respectively). The nucleocapsid protein
of MoMuLV (NCp10) facilitates hybridization of tRNAPro
with viral RNA by
preferentially binding to T1, T2, T3 and V1 sites consisting of four
nucleotides (Miller et al. 2014).
2.2.2. Formation of the template–primer duplex
In order to serve as a primer, the 18 nucleotides at the 3' end of the tRNA
must pair with the 18 nucleotides of the PBS sequence located in the 5'
region of the viral genome (see Figure 2.2). In the gRNA and the tRNA
primer, the 18 nucleotides are not accessible, as several of them are involved
in intramolecular base-pairing interactions forming part of the secondary
structure of these molecules (see Figure 2.3). These interactions must
therefore be broken to form the intermolecular base-pairing interactions
leading to the binding of the tRNA primer to the PBS sequence of the
gRNA. For this reason, it is necessary to use a temperature of 70°C to
reconstitute the HIV-1 and MoMuLV tRNA–gRNA hybrids in vitro in the
absence of protein (Isel et al. 1993; Fossé et al. 1998). In contrast, it is
possible to reconstitute these hybrids in vitro at physiological temperature
when the reaction medium contains the NC of MoMuLV, RSV or HIV-1
(Prats et al. 1988; Barat et al. 1989). These latter results led to the proposal
that NC is the viral protein largely responsible for tRNA primer binding to
the PBS sequence. In the case of HIV-1, human RNA helicase A could
facilitate hybridization by inducing a conformational change in the 5'-UTR
region of the gRNA (Xing et al. 2011, 2012).
Nuclear magnetic resonance (NMR) analysis of the complexes that the
MoMuLV NC forms with tRNAPro
and the PBS region has determined the
mechanism of formation of the tRNAPro
–PBS region duplex in MoMuLV
(Miller et al. 2014). Upon binding to the V1 site of the viral RNA and the T1
site of the tRNAPro
(see Figure 2.3), the NC releases three nucleotides from
the PBS sequence and also three nucleotides from the anti-PBS sequence of
the tRNAPro
to initiate the base-pairing interactions of the primer–template
duplex. In addition, NC binding to the T2 and T3 sites destabilizes the

three-dimensional structure of the tRNAPro
, and thus increases the
accessibility of the anti-PBS sequence. The hybridization of tRNALys3
to the
HIV-1 PBS sequence has also been studied by NMR (Tisné et al. 2004;
Barraud et al. 2007). These studies indicate that in the anti-PBS sequence of
tRNALys3
, a few nucleotides in the T arm and the 3' CCA end initiate the
formation of the primer–template duplex by base-pairing. This initiation
process does not require the presence of the NC, but it accelerates it. In
contrast, the formation of the long PBS-anti-PBS duplex (18 base pairs)
requires the action of the NC which destabilizes the three-dimensional
structure of the tRNALys3
by binding to the D-arm loop.
The distinct roles of the Gag precursor and NC on tRNA hybridization to
the PBS sequence have primarily been studied in HIV-1 (Feng et al. 1999;
Guo et al. 2009; Seif et al. 2015; Jin and Musier-Forsyth 2019). The
secondary structures of the hybrid within mature and immature viral
particles can be studied using the SHAPE method (Wilkinson et al. 2008;
Seif et al. 2015). From these studies, a two-step hybridization model was
proposed (see Figure 2.4):
– the formation of an unstable duplex, which is facilitated by the NC
domain of the Gag precursor prior to its proteolysis during the maturation of
the virus particle;
– the NC in the mature virus particle facilitates the conversion of the
unstable duplex to a stable duplex.
The secondary structure model of the stable duplex shows that the
tRNALys3
–gRNA interaction is not limited to the PBS sequence and also
involves two additional interactions consisting of short sequences (four and
six nucleotides). Interestingly, the hybrid reconstituted in vitro at 70°C in the
absence of protein can mirror that present in the virion. Indeed, under these
conditions, all three interactions between tRNALys3
and gRNA have also
been observed (Isel et al. 1995). It should be noted, however, that the two
additional interactions would only occur in some HIV-1 isolates (Sleiman
et al. 2012). In the case of RSV, the tRNATrp
–gRNA interaction would not
be limited to the PBS sequence but would also involve another sequence
consisting of seven nucleotides (Aiyar et al. 1992; Morris et al. 2002). In
contrast, the tRNAPro
–gRNA interaction is limited to the PBS sequence in
MoMuLV (see Figure 2.5) (Fossé et al. 1998).

Figure 2.4. Secondary structures of the HIV-1 tRNA
Lys3
–gRNA hybrid. For a
COMMENTARY ON FIGURE 2.4. – In HIV-1, the Gag precursor facilitates the
formation of an unstable hybrid between the tRNALys3
and the PBS sequence
(secondary structure on the left). The NC generated by Gag maturation
converts the unstable hybrid to a stable hybrid (Seif et al. 2015). The PBS
sequence is only partially paired with the 3' end of the tRNALys3
in the
unstable hybrid. The tRNALys3
is in purple; the nucleotides shown are mainly
those that pair with the gRNA, while the others are replaced by the thick
line. The PBS sequence is in blue, while the other two sequences of the
gRNA that interact with the tRNALys3
are in green. The first three nucleotides
to be copied by RT are in orange.
Figure 2.5. Secondary structure of the MoMuLV–gRNA
Pro
hybrid. For a
V2 site

COMMENTARY ON FIGURE 2.5. – The PBS sequence of the MoMuLV gRNA is
fully paired with the 3' end of the tRNAPro
. The tRNAPro
is in purple; the
nucleotides shown are mainly those that pair with the gRNA, while the
others are replaced by the thick line. The PBS sequence is in blue. The first
three nucleotides to be copied by RT are in orange. By binding to the V2 site,
NC destabilizes the apical stem and thus facilitates extension of the tRNAPro
primer by RT.
2.2.3. Role of the structure of the template–primer duplex upon
initiation
The determination of the secondary structure of the HIV-1 and MoMuLV
duplexes (Isel et al. 1995; Fossé et al. 1998), using chemical and enzymatic
probes, has shown that the first three nucleotides to be copied by RT are
unpaired and adjacent to a stem-loop (see Figures 2.4 and 2.5). Binding of
the MoMuLV NC to the apical loop corresponding to the V2 site (see
Figure 2.5) facilitates extension of the tRNAPro
primer by RT by
destabilizing the apical stem of the viral RNA (Miller et al. 2014).
The reverse transcription initiation step corresponding to a short primer
extension has been studied in detail in HIV-1 (Isel et al. 2010; Sleiman et al.
2012). This step is characterized by RT pauses during primer extension,
rapid dissociation of RT from the primer–template duplex and a DNA
polymerization rate that is slow. RT pausing leads to the accumulation of
two intermediates corresponding to the addition of three and five nucleotides
to the 3’ end of the tRNALys3
primer, respectively. The elongation step of
reverse transcription, which is fast and processive, starts as soon as the sixth
nucleotide is added to the elongated primer.
The interaction of RT with the primer–template duplex during initiation
can be characterized at the molecular level by a single-molecule study based
on fluorescence resonance energy transfer (FRET) (Liu et al. 2010). This
study shows that RT can bind to the primer–template duplex in two opposite
orientations: one competent for DNA synthesis because the polymerase site
of the enzyme is at the 3' end of the primer and one inactive for DNA
synthesis because the polymerase site is opposite to the 3’ end of the primer
(see Figure 2.6). In the absence of deoxyribonucleotides, both binding modes
are observed.

P
H
C
A
G
5’
3’
5’
H
C
A
G
5’
3’
5’
P
H
U
A
G
C
A
G
5’
3’
5’
P
G
T
C
A
T
C
G
T
C
0
+3
+6
Figure
2.6.
HIV-1
RT
orientations
on
the
primer–template
duplex.
For
a
color
version
of
this
figure,
see
www.iste.co.uk/fosse/structures.zip

COMMENTARY ON FIGURE 2.6. – On the primer–template duplex consisting
of the gRNA and tRNALys3
PBS sequence, the HIV-1 RT can adopt two
opposite orientations in the absence of deoxyribonucleotides (state 0). In the
diagram, only one orientation is shown in state 0, where the RT polymerase
site (P) is near the 3' end of the tRNALys3
(purple molecule). After adding
three (state +3) or five (state +5 not shown here) deoxyribonucleotides, the
RT adopts an orientation that is not competent for DNA polymerization,
because it is its RNase H (H) site that is close to the 3' end of the primer
extended by three or five nucleotides. By adding the sixth nucleotide (+6
state), the RT adopts the competent orientation for DNA synthesis and thus
begins to start the elongation step of reverse transcription, which is coupled
with the destabilization of the stem-loop template (blue molecule).
In the presence of deoxyribonucleotides, the orientation competent for
DNA synthesis is promoted until three nucleotides are incorporated at the
end of the primer. This three-nucleotide extension increases the likelihood of
RT pausing because it allows the stem-loop located upstream of the primer
to impose the inactive orientation of the enzyme for DNA polymerization.
This orientation is prevalent until the incorporation of the fifth nucleotide.
Incorporation of the sixth nucleotide destabilizes the stem-loop, and thus
allows the RT to adopt the orientation competent for polymerization (see
Figure 2.6). The destabilization of the stem-loop is thought to be facilitated
by the NC (Wilkinson et al. 2008; Liu et al. 2010).
2.3. RNA structures involved in the first strand transfer
2.3.1. Actors involved in the first strand transfer
The first strand transfer, which is an essential step in reverse
transcription, occurs at the R repeat sequence that is present at both ends of
the gRNA (see Figure 2.2, step 2). The length of the R sequence varies by
retroviral species (e.g. 16 nucleotides in mouse mammary tumor virus
(MMTV) and 247 nucleotides with human T-cell leukemia virus type 2
(HTLV-2)) (René et al. 2018). The transfer primarily corresponds to a
base-pairing interaction between the R region located at the 3' end of the
gRNA and the r region of the newly synthesized DNA strand, called
strong-stop DNA (ssDNA). The ssDNA is synthesized by the RT which
copies the RU5 region of the gRNA. The first strand transfer thus requires at
least three players: gRNA, RT and ssDNA. RNase H activity of RT, leading

to degradation of the RU5 RNA region that has been copied, is required for
the transfer reaction because it releases the ssDNA (Tanese et al. 1991;
Peliska and Benkovic 1992; Blain and Goff 1995; Chen et al. 2003).
Simplified experimental systems using RT, two short RNAs synthesized in
vitro and a DNA primer have been able to efficiently reproduce the first
strand transfer in vitro when the R region is truncated (Peliska and Benkovic
1992; Werner et al. 2001). In contrast, the experimental system must contain
the NC to obtain in vitro efficient transfer when both RNAs are relatively
long and contain an entire R sequence (Darlix et al. 1993; Allain et al. 1994;
Guo et al. 1997). Under these conditions, NC increases the transfer by
five- to tenfold (Guo et al. 1997; Chen et al. 2003). In addition to RT, NC
thus appears to be another viral protein playing an important role in the first
strand transfer.
2.3.2. Molecular basis of R–r pairing
The molecular mechanisms governing R–r pairing between the 3' end of
gRNA and the 3' end of ssDNA have been primarily studied in HIV-1. In
vitro analysis of ssDNA with a chemical probe and two enzyme probes
suggests that NC binding to four preferential sites triggers unfolding of the
ssDNA three-dimensional structure and thus facilitates the interaction of the
r-sequence with the gRNA R-sequence (Chen et al. 2016b). An in vitro study
(Kanevsky et al. 2005) and an ex vivo study using SHAPE (Watts et al.
2009) have shown that the 3' R sequence of the gRNA folds to form two
stem-loops called TAR and poly(A). It has also been shown in vitro (Chen
et al. 2016b) that the r-sequence of the ssDNA forms the cTAR and cpoly(A)
stem-loops that are complementary to the TAR and poly(A) stem-loops,
respectively (see Figure 2.7). The unfolding of the secondary structures
constituted by these stem-loops is also necessary to allow r–R pairing
corresponding to a stable duplex of 95 base pairs.
In vitro and ex vivo studies suggest that TAR/cTAR stem-loops play a
more important role in r–R pairing than poly(A)/cpoly(A) stem-loops (Ohi
and Clever 2000; Berkhout et al. 2001; Moumen et al. 2001). The interaction
of NC with the TAR RNA stem-loop can be characterized in vitro at the
molecular level by a single-molecule study using optical tweezers
(McCauley et al. 2015) and by an NMR study (Belfetmi et al. 2016). The
single-molecule study showed that NC can destabilize the TAR stem-loop by

targeting four paired guanines that are adjacent to low-stability regions
(bulges and G-U pair) (see Figure 2.8a).
NMR has shown that among these guanines, the one located in the apical
stem plays the most important role, as it corresponds to the site where the
NC initiates the destabilization of the TAR stem-loop.
Biochemical and biophysical studies performed in vitro with the cTAR
stem-loop in the absence of NC (Bernacchi et al. 2002; Azoulay et al. 2003;
Vo et al. 2009; Kanevsky et al. 2011) and ssDNA (Chen et al. 2016b)
revealed a dynamic structure for this stem-loop, which results from the
equilibrium between two conformations (see Figure 2.8b).
The same studies have shown that the NC shifts the equilibrium to the
more open conformation (see Figure 2.8b).
Figure 2.7. Secondary structures of the 3' ends of gRNA and ssDNA. Secondary
structures were determined using chemical and enzymatic probes with the
HIV-1 MAL isolate (Kanevsky et al. 2005; Chen et al. 2016b). The complementary r
and R sequences are framed by the dashed lines. For a color version of this figure,
see www.iste.co.uk/fosse/structures.zip

a)
b)
Figure 2.8. a) Effect of NC on the TAR stem-loop; b) effect of NC on the cTAR
stem-loop. For a color version of this figure, see www.iste.co.uk/fosse/structures.zip
COMMENTARY ON FIGURE 2.8. – NC destabilizes the TAR RNA stem-loop
shown in panel A by preferentially interacting with the four guanines
colored in orange and red. The destabilized G–C base pairs are outlined by

a dashed line. Destabilization is initiated at the red-colored guanine. The NC
destabilizes the cTAR DNA stem-loop presented in panel B by destabilizing
four base pairs, thus enabling the opening of the bottom of the stem.
Figure 2.9. Initiation models of r–R pairing via TAR and cTAR. TAR–cTAR duplex
formation is initiated either from the bottom of stem-loops that are destabilized by NC
(zipper pathway) or from apical loops that are accessible (kissing pathway). The
initial interactions limited to a few base pairs require the action of the NC to expand
and form the stable TAR–cTAR duplex. For a color version of this figure, see
Based on the studies mentioned above and performed with isolated TAR
and cTAR stem-loops, two models of r–R pairing initiation have been proposed
(see Figure 2.9). In both models, NC is absolutely required to convert the
initiation complexes into the TAR–cTAR duplex. In one model, initiation
occurs at the bottom of the stems (zipper pathway), whereas in the other model,
it occurs at the apical loops (kissing pathway). One of the two initiation

pathways is not favored when r–R pairing is performed in vitro with the whole
ssDNA and the 3' end of the gRNA; a third pathway is thought to involve the
poly(A) and cpoly(A) stem-loops (Chen et al. 2016b). There are no studies that
have been able to determine how ex vivo/in vivo r–R pairing is initiated.
2.4. RNA structures promoting genetic recombination
Retroviral recombination is implicated in retrovirus escape from the host
immune system and the development of multiple resistance to antiretroviral
agents (Simon-Loriere et al. 2011). There are between 5 and 14 recombination
events per genome during an HIV-1 replicative cycle (Cromer et al. 2016).
These events occur when two gRNA molecules that are genetically different
but similar have been packaged into the same viral particle as a heterodimer.
Retroviral recombination depends primarily on the internal strand transfer that
occurs during reverse transcription; more precisely, the synthesis of the
negative polarity DNA strand (see Figure 2.2, step 3). A recombination event
is the result of an internal strand transfer event.
2.4.1. Internal strand transfer
The molecular mechanisms governing internal strand transfer are similar
to those involved in the first strand transfer. Specifically, the RNA template,
corresponding to one strand of the heterodimer, must be degraded by the
RNase H activity of RT after being copied by this enzyme in order to allow
the release of the newly synthesized DNA and its pairing with the acceptor
RNA corresponding to the other strand of the heterodimer (see Figure 2.10).
Five to fourteen strand transfer events can occur along the gRNA during an
HIV-1 replication cycle, because the two strands of gRNA constituting the
heterodimer are similar and therefore contain long identical regions where
base-pairing of the newly synthesized DNA with the acceptor RNA is
possible. Internal strand transfer has been studied mainly in vitro, as it is
difficult to characterize ex vivo/in vivo molecular mechanisms (Basu et al.
2008). Simplified experimental systems using RT, two in vitro synthesized
RNAs and a DNA primer have been used to reproduce in vitro internal
strand transfer (Derebail et al. 2003). Under these experimental conditions,
NC strongly increases internal strand transfer in regions where the RNA is
highly structured. In contrast, the stimulatory effect of NC on internal strand
transfer is low in weakly structured RNA regions.

Figure
2.10.
Internal
strand
transfer.
For
a
color
version
of
this
figure,
see

COMMENTARY ON FIGURE 2.10. – Example of the acquisition of resistance to
two antiretrovirals by internal strand transfer. The gRNA dimer consisting of
two RNA strands of positive polarity is shown as two lines colored purple
(acceptor RNA) and blue (donor RNA). The region that associates the two
subunits of the dimer is shown by the two juxtaposed lines. The red star on
the donor RNA corresponds to a mutation conferring resistance to an
antiretroviral targeting RT, and the one on the acceptor RNA corresponds to
a mutation conferring resistance to an antiretroviral targeting the viral
protease. The DNA strand of negative polarity being synthesized is
represented by the brown arrow. The synthesis of this strand starts on the
donor RNA strand and thus contains the copy of the mutation conferring
resistance to an antiretroviral targeting RT. (1) After degradation by the
RNase H activity of the RT of the part of the RNA that has just been copied
(dashed line), internal strand transfer occurs by pairing of the newly
synthesized DNA with the complementary part of the acceptor RNA. (2) DNA
strand synthesis continues, and the strand contains a copy of the mutation
conferring resistance to a protease-targeting antiretroviral. (3) Subsequent
steps in reverse transcription lead to double-stranded viral DNA containing
both mutations.
2.4.2. RNA structures triggering internal strand transfer
Several in vitro studies have shown that the structure of the acceptor
RNA plays an important role in internal strand transfer (Negroni and Buc
2000; Moumen et al. 2003; Hanson et al. 2005). The transfer reaction
depends generally on a stem-loop present in the acceptor RNA (Moumen
et al. 2003; Galetto et al. 2004). Depending on the acceptor RNA, the crossover
point, that is, the site where the RT starts to copy the acceptor RNA, is
located in or near the stem-loop (Roda et al. 2002; Balakrishnan et al. 2003;
Moumen et al. 2003). Depending on the template RNA, the NC changes
(Negroni and Buc 2000; Hanson et al. 2005) or does not change (Roda et al.
2002; Balakrishnan et al. 2003) the crossover point. Internal strand transfer
is also facilitated by RNA structures resulting from interactions between the
donor and acceptor strands. These structures bringing the two strands
together can be two stem-loops that interact via their apical loop or tetrads of
guanines (Balakrishnan et al. 2003; Shen et al. 2011).
The role of acceptor RNA structure in internal strand transfer is
incorporated into the acceptor invasion model (Negroni and Buc 2000) that

is consistent with the results of many studies (see references cited by Basu
et al. (2008)). In this model (see Figure 2.11), degradation of a portion of the
donor RNA by the RNase H activity of RT creates an invasion site where a
single-stranded DNA region can interact with the acceptor RNA. This
invasion step corresponds to the first base-pairing interaction between the
newly synthesized DNA and the acceptor RNA that occurs upstream of the
crossover point. This initial interaction expands by displacing the donor
RNA fragments, which had remained paired to the DNA. The template
exchange occurs during DNA polymerization when the 3' end of DNA pairs
with the acceptor RNA. Thanks to its nucleic acid chaperone activity, NC
facilitates the above-mentioned molecular mechanisms and thus stimulates
internal strand transfer.
Figure 2.11. Acceptor invasion model. For a color version
2
1
Donneur 5’ (+)
5’ (-)
3’
Accepteur 5’ (+) 3’
3
4
5
Donor
Acceptor
5’ (+)
5’ (-)
3’
3’
5’ (+)

COMMENTARY ON FIGURE 2.11. – Internal strand transfer is triggered by the
stem-loop present in the acceptor RNA. The acceptor and donor RNA
strands are in blue and purple, respectively. Dashed lines indicate the
regions degraded by RT RNase H activity. The negative polarity DNA being
synthesized is represented by the brown arrow. Step 1, the RT copies the
donor strand and degrades the copied part through its RNase H activity.
Step 2, in this example, the acceptor invasion site corresponds to the apical
loop, the nucleotides of which are accessible and can pair with the
complementary part of the newly synthesized DNA that is released from the
donor strand by the RNase H activity of the RT and the NC. Step 3,
the initial DNA–acceptor RNA interaction expands and is facilitated by the
NC while the RT continues to copy the donor RNA. Step 4, the 3' end of the
DNA pairs with the complementary portion of the acceptor strand. Step 5,
DNA synthesis continues on the acceptor strand, the copied portion of which
is in turn degraded by the RNase H activity of the RT.

3
RNA Structures Regulating the
Expression of the Retroviral Genome
3.1. Regulatory RNA structures of proviral DNA transcription
3.1.1. The transcriptional activator Tat
Among the different retroviral species (see Table 1.1), those belonging to
the genus lentivirus are characterized by, among other things, their ability to
produce the regulatory protein transactivator of transcription (Tat), which is a
transcriptional activator. The activation of the transcription of proviral DNA
by Tat is, depending on the lentivirus, independent of or dependent on an
RNA structure called the transactivator response element (TAR). The
lentiviruses bovine immunodeficiency virus (BIV), equine infectious anemia
virus (EIAV), Jembrana disease virus (JDV), simian immunodeficiency virus
(SIV), HIV-1 and HIV-2 belong to the group where transcriptional activation
is dependent on Tat binding to TAR (Ott et al. 2011).
Despite their small size (75–101 amino acids), Tat proteins, interacting
with TAR structures, have two essential domains that are conserved among
different lentiviruses (see Figure 3.1). The transactivation domain is
cysteine-rich and has a highly conserved motif called the core. This motif
plays a crucial role in the transactivating function of Tat proteins, as it is
directly involved in the interaction with the positive elongation factor
P-TEFb. The RNA-binding domain contains an arginine-rich basic motif,
which confers its function. In addition, this motif plays an important role
in Tat’s stability and nuclear localization. Tat proteins are inherently
unstructured and structure themselves by interacting with their ligands (Ott
et al. 2011; Schulze-Gahmen and Hurley 2018).

Figure
3.1.
Alignment
of
six
Tat
protein
sequences.
For
a
color
version
of
this
figure,
see

COMMENTARY ON FIGURE 3.1.– The numbers in parentheses indicate the
numbers of amino acids that are present in the N- and C-terminal ends but
not represented in this alignment. Above the HIV-1 Tat sequence are the
amino acid positions. Conserved cysteines and histidines are highlighted in
yellow. Conserved core amino acids are highlighted in purple. The
arginine-rich motif is highlighted in blue. TD indicates the transactivation
domain. RBD indicates the RNA binding domain.
In lentiviruses whose activation of transcription depends on the Tat–TAR
interaction, the molecular mechanisms governing the latter are similar but
not identical. In all of them, the TAR structure is formed by a part of the R
sequence located at the 5' end of the genome, but it can be formed by one or
two stem-loops.
3.1.2. TAR structures with one stem-loop
BIV, EIAV, JDV and HIV-1 lentiviruses possess a single stem-loop TAR
structure (Feng and Holland 1988; Carpenter et al. 1993; Chen et al. 1999;
Anand et al. 2008). With the exception of EIAV, a bulge composed of one to
three nucleotides plays an essential role in Tat protein binding to TAR RNA.
The TAR stem-loop is relatively long in HIV-1 (between 53 and 55
nucleotides depending on the different isolates), whereas it is rather short
(between 25 and 28 nucleotides) in BIV, EIAV and JDV lentiviruses (see
Figure 3.2). The apical loop and the bulge of the HIV-1 TAR stem-loop are
essential for the activation of transcription. In contrast, the lower part of the
stem-loop is not required.
3.1.2.1. TAR–Tat interaction in HIV-1
The activation of proviral DNA transcription by the Tat–TAR interaction
has been most extensively studied in HIV-1. In the absence of this
interaction, HIV-1 genome expression can only be very weak and produces
mainly short RNAs resulting from transcripts that are stopped at the
elongation step (Ott et al. 2011). A positive feedback loop regulates Tat
protein production. Specifically, the initial synthesis of Tat resulting from
very low-level transcription in the absence of this protein will allow the
protein to activate transcription of the viral genome, and thus iteratively
increase Tat production and viral particle production (Ott et al. 2011).
RNA Structures Regulating the Expression of the Retroviral Genome 33

Figure 3.2. TAR structures with one stem-loop
COMMENTARY ON FIGURE 3.2. – Each TAR stem-loop is formed by the 5' end
of the R sequence of the viral genome. The HIV-1 TAR stem-loop shown
corresponds to that of the NL4-3 isolate with a three-nucleotide bulge; in
some isolates, the bulge is two nucleotides. Although the HIV-1 TAR
stem-loop is much longer than other stem-loops, it is only the top half (boxed
portion) that is involved in transcription activation. The asterisk denotes the
guanine that interacts with the Tat protein loop in the Tat–SEC–TAR
complex.
In the absence of Tat, and in the majority of cases, the transcription of
proviral DNA is not continued beyond the initiation step (see Figure 3.3).
The elongation step of transcription depends on Tat’s recognition of the
bulge formed by the TAR stem-loop. The arginine-rich motif of Tat
interacts with the nucleotides constituting the bulge, and also those nearby
that form the major groove of the RNA double helix (Schulze-Gahmen and
Hurley 2018; Chavali et al. 2019). The affinity of Tat for the TAR stem-loop
is low and the Tat–TAR interaction is not sufficient to activate proviral DNA

transcription, which is a complex process. Indeed, Tat must recruit the super
elongation complex (SEC) consisting of the transcription factors ENL/AF9
and ELL2, AFF4 protein, cyclin T1 and CDK9 kinase (Schulze-Gahmen et
al. 2016). Cyclin T1 and Tat each interact with AFF4 (see Figure 3.3). The
transactivation domain of Tat (see Figure 3.1) directly interacts with cyclin
T1. The positive elongation factor P-TEFb resulting from the cyclin
T1–CDK9 association inherently lacks the ability to bind to TAR (Romani
et al. 2010). In contrast, the affinity of the Tat–P-TEFb complex for TAR is
higher than that of Tat alone because cyclin T1 interacts with the apical
loop (see Figure 3.3). In addition, the Tat–P-TEFb complex has its
affinity increased for the TAR stem-loop when associated with AFF4
(Schulze-Gahmen et al. 2016). In the Tat–SEC–TAR complex, the Tat loop,
which is composed of amino acids N24–Y26 (see Figure 3.1) and results
from zinc coordination by cysteines, directly interacts via hydrogen bonds
with a guanine present in the apical loop of the TAR RNA (see Figure 3.2)
(Schulze-Gahmen and Hurley 2018). Formation of the Tat–SEC–TAR
complex stimulates the elongation step of transcription by allowing CDK9 to
phosphorylate the C-terminus of RNA polymerase II and the negative
elongation factors NELF and DSIF.
3.1.2.2. TAR–Tat interaction in SIVcpz, BIV, JDV and EIAV
lentiviruses
The activation of transcription by TAR–Tat interaction in SIVcpz (SIV
infecting chimpanzees) is similar to that observed in HIV-1 (D’Orso and
Frankel 2009). In contrast, in bovine lentiviruses (BIV and JDV), the
activation of transcription is different because P-TEFb has the primary role
of phosphorylating the c-terminus of RNA polymerase II. More specifically,
cyclin T1 does not interact with the TAR stem-loop, and Tat has a high
affinity for it that is independent of its interaction with cyclin T1 (D’Orso
and Frankel 2009). Tat does not interact with the apical loop of TAR but
only with the major RNA groove located near the central bulge,
corresponding to UGU in BIV and UA in JDV (see Figure 3.2).
In EIAV, the activation of transcription also requires the formation of the
P-TEFb–Tat complex, which depends on the insertion of the C-terminal
leucine of Tat (see Figure 3.1) into the hydrophobic pocket formed by two
helices of cyclin T1 (Anand et al. 2008). The arginine-rich motif of Tat and
adjacent amino acids interact with the apical loop and the upper part of the

TAR stem-loop. Cyclin T1 increases the affinity of the P-TEFb–Tat complex
for TAR because it interacts with part of the apical loop (Anand et al. 2008).
Figure 3.3. Activation of transcription by the HIV-1 Tat protein. For a
COMMENT ON FIGURE 3.3.– In the absence of Tat, cellular transcription
factors (e.g. NF-κB and Sp1) allow the initiation of transcription by RNA
polymerase II by binding to the U3 portion of the 5' LTR. This initiation step,
which is not followed by an elongation step (shown by an arrow with a red
cross), generates short transcripts (50–60 nucleotides) that can form the
upper part of the TAR stem-loop. Elongation by the polymerase is inhibited
by the protein factors NELF and DSIF. The elongation step requires that Tat
recruits the super elongation complex (SEC) which consists of the proteins

cyclin T1 (CycT1), CDK9, AFF4, ENL/AF9 and ELL2. Tat and CycT1 are
the two proteins in the Tat–SEC complex that bind it to TAR RNA. Tat
interacts with the bulge and part of the apical loop, whereas CycT1 interacts
with the apical loop. The Tat–SEC–TAR interaction leads CDK9 to
phosphorylate NELF and DSIF and thus counteract their inhibitory effect on
elongation. CDK9 hyperphosphorylation of the C-terminus of RNA
polymerase II leads the latter to perform elongation.
3.1.3. TAR structures with two stem-loops
SIVmac (SIV infecting macaques) and HIV-2 are representative of
lentiviruses with a two stem-loop TAR structure (D’Orso and Frankel 2009)
(see Figure 3.4). Each stem-loop has a central two-nucleotide bulge (UU and
UA). The stem-loop with the UU bulge appears to play a more important
role than the other stem-loop in activating transcription (Fenrick et al. 1989;
Rhim and Rice 1994). Transcription activation also depends on the
formation of the P-TEFb–Tat complex (D’Orso and Frankel 2009).
Figure 3.4. TAR structure of SIVmac and HIV-2 lentiviruses. The two-stem-loop
TAR structure of SIVmac is identical to that of HIV-2 because the part of the R
sequence that adopts this structure is the same in these two lentiviruses
Each of the two stem-loops of the TAR structure binds a P-TEFb–Tat
complex via an interaction involving Tat and the central bulge. Unlike
HIV-1, extensive structural studies of P-TEFb–Tat–TAR complexes have

not been performed with Tat proteins and TAR RNAs from SIVmac and
HIV-2 lentiviruses.
3.2. RNA structures regulating genomic RNA maturation
Figure 3.5. Maturation of retroviral RNA. For a color version
COMMENTARY ON FIGURE 3.5.– 1) Proviral DNA is transcribed by RNA
polymerase II. 2) Primary structure of the pre-messenger RNA resulting
from transcription and initial post-transcriptional modifications: addition of
a 5' cap, cleavage of the 3' end and synthesis of a poly(A) tail. 3) Splicing: a
single event producing, for example, the env mRNA in ALV alpharetrovirus
or three events producing, for example, the nef mRNA in HIV-1 lentivirus.
The start of transcription is indicated by a horizontal black arrow. The
orange arrow indicates the 3' cleavage. The orange disk represents the cap.
Retroviral RNAs, like all cellular pre-messenger RNAs synthesized by
RNA polymerase II, are the products of rapid maturation leading to the
gag pol env
5’ LTR 3’ LTR
U3 R U5
U3 R U5
1
2
3
AAAAAA 3’
5’ 3’
5’
5’ AAAAAA 3’
5’ AAAAAA 3’

addition of a cap at their 5' end and cleavage of their 3' end followed by the
addition of a poly(A) tail consisting of approximately 200 adenosine
monophosphate residues (see Figure 3.5). Maturation continues with one or
two splicing events in simple retroviruses (e.g. ALV and RSV
alpharetroviruses) and multiple splicing events in complex retroviruses (e.g.
HIV-1 and HIV-2 lentiviruses). A characteristic of retroviruses is that the
majority of capped and polyadenylated RNA molecules are not spliced
(McNally 2008). In effect, the unspliced RNA serves as single-stranded
genomic RNA which is converted to double-stranded DNA by RT or as
mRNA for the synthesis of Gag and Gag–Pol polypeptide precursors.
Retroviruses have therefore evolved strategies to control the proportions of
spliced and unspliced mRNAs. Cis-regulatory elements are involved in this
control (Cochrane et al. 2006). The cis-regulatory properties of some are
related to the RNA structures they form. These structures have only been
well-characterized in RSV and HIV-1.
3.2.1. The negative regulator of splicing of RSV
The RSV belonging to the genus alpharetrovirus was one of the first
retroviruses to be studied because, in addition to the gag, pol and env genes,
it contains the oncogene sarc (src) which codes for a tyrosine kinase
involved in cell proliferation. In addition, this virus has served as a relatively
simple model for studying alternative splicing in retroviruses. Indeed, the
unspliced mRNA representing approximately 75% of RSV mRNAs (Hudson
et al. 2016) contains a single splice donor site and two acceptor sites to
generate env and src mRNAs (see Figure 3.6).
The low proportion of spliced RNAs (approximately 25%) is primarily
due to three negative splicing signals (Cochrane et al. 2006). The first results
from the env mRNA branch point, which only partially matches the
canonical sequence required for efficient splicing (Katz and Skalka 1990).
The second is due to degeneration of the polypyrimidine motif constituting
part of the acceptor site of the src mRNA (Zhang and Stoltzfus 1995). The
third corresponds to the negative regulator of splicing (NRS) that reduces the
production of env and src mRNAs (Arrigo and Beemon 1988; Cochrane et
al. 2006). This cis-regulatory element also has a very long-range stimulatory
effect on polyadenylation of unspliced mRNA (Hudson et al. 2016). The
NRS consists of approximately 230 nucleotides and is located in the gag
gene (see Figure 3.6).

Figure 3.6. RSV unspliced mRNA. The RSV unspliced mRNA contains one splice
donor site (SD site) and two acceptor sites (SA sites) that generate the env and src
mRNAs. The NRS sequence, which is colored green, is more than 4,000 nucleotides
from the first SA. The orange disk represents the 5' cap. Regions and genes are not
shown to scale. For a color version of this figure, see www.iste.co.uk/fosse/
structures.zip
The secondary structure of NRS was determined using chemical and
enzymatic probes (Bar et al. 2011). The 5' and 3' ends of the NRS interact
to form a cruciform structure that is conserved among different
alpharetroviruses (see Figure 3.7).
Figure 3.7. Cruciform structure of the NRS. For a color version

COMMENTARY ON FIGURE 3.7.– The 5' and 3' ends of NRS interact to form a
cruciform structure that is responsible for the splicing inhibitory activity of
NRS. Helices 1 and 2 are called H1 and H2, respectively. Stem-loops 1 and
2 are called SL1 and SL2, respectively. The apical part of SL2 partially
corresponds to the splice donor site consensus sequence (AGGURAGU, R
being a purine). The orange-colored nucleotides are complementary to the
U1 small RNA region that matches the splice donor consensus sequence.
The cruciform structure is responsible for the splicing inhibitory effect of
NRS. The hnRNP H and SR proteins (ASF/SF2, 9G8 and SC35), which are
splicing-regulating factors, bind to the cruciform structure. The splicing
inhibitory activity of NRS has not been shown to be dependent on the
binding of hnRNP H proteins. Stem-loop 1 is the major binding site for the
SR proteins ASF/SF2 and 9G8 (McNally and McNally 1996; Bar et al.
2011). The H2 helix also binds the SR 9G8 protein, whereas the H1 helix
and stem-loop 2 are not binding sites for this protein. The activity of NRS is
related, at least in part, to its interaction with the SR 9G8 protein. The results
obtained with SR proteins led to the hypothesis that SR proteins recruit the
nuclear ribonucleoprotein U1, which recognizes a pseudo-splice donor site
in NRS. Indeed, the apical part of the stem-loop 2 has a pseudo-donor site
that enables the binding of the small nuclear RNA U1 that participates in
splicing (see Figure 3.7). The conformation of stem-loop 2 must not be
altered in order to bind the U1 RNA (Cabello-Villegas et al. 2004). This
binding leads to the formation of a non-functional spliceosome, in which the
pseudo-donor site is coupled with one of the 3' acceptor sites. This coupling
prevents the 3' acceptor site from interacting with the authentic 5' donor site.
It has also been hypothesized that the SR proteins are required for the
coupling of the pseudo-donor site to one of the 3' acceptor sites.
Furthermore, by binding to the NRS, SR proteins allow the NRS to stimulate
3' polyadenylation of the newly synthesized RNA (Hudson et al. 2016).
3.2.2. Structural diversity and alternative splicing in HIV-1
HIV-1 pre-messenger RNA (~9,200 nucleotides) undergoes extensive
alternative splicing that produces more than 50 mRNA isoforms, allowing for
optimized expression of viral proteins required for viral replication (Sertznig
et al. 2018). This is due to the presence of at least four donor sites and eight

acceptor sites in all HIV-1 isolates. The many possibilities of alternative
splicing were illustrated by the identification of 109 spliced RNAs when
analyzing a clinical HIV-1 isolate with massive parallel sequencing (Ocwieja
et al. 2012). In this particular case, the pre-messenger RNA contains 7 donor
sites and 14 acceptor sites. Alternative splicing results from various complex
mechanisms. One of these mechanisms relies on the strength of a donor or
acceptor site, which depends on the degree of similarity to the consensus
sequence (Sertznig et al. 2018). The greater the difference between the site
sequence and the consensus sequence, the lower its strength, as well as the
amount of mRNA resulting from splicing at that site. Another mechanism
involves cis-regulatory elements that, by binding proteins, activate or inhibit
splicing at a given site. Splicing at a site may depend on a combination of
these two mechanisms.
A third recently discovered mechanism relies on the regulation of
alternative splicing by the structural diversity of HIV-1 pre-messenger RNA.
Indeed, pre-messenger RNA does not adopt a single conformation but
several alternative conformations ex vivo (Tomezsko et al. 2020).
Pre-messenger RNA molecules are therefore present in different
conformations in the nucleus of the infected cell. Depending on the
conformation adopted by a pre-messenger RNA molecule, splicing at a given
site can be repressed or activated. The structural diversity of HIV-1
pre-messenger RNA partly explains its alternative splicing and the
persistence of unspliced pre-messenger RNA, which acts both as genomic
RNA and as mRNA for the gag and pol genes. The role of RNA
conformation was best illustrated by studying the A3 acceptor site, which is
required for the production of Tat mRNAs. The regulation of splicing by the
RNA structure at the A3 site can reduce the amount of Tat mRNAs by a
factor of 100 (Tomezsko et al. 2020). Specifically, the region of the
pre-messenger RNA containing the A3 site forms two distinct and major
secondary structures (see Figure 3.8). In one (structure 1), present in 63–67%
of pre-messenger RNA molecules, the pyrimidine-rich sequence of the A3
site is fully accessible and can therefore bind the U2AF protein that plays a
crucial role in 3' splicing. In contrast, in the other secondary structure
(structure 2), present in 33–37% of pre-messenger RNA molecules, this
sequence is inaccessible to U2AF because it is largely engaged in
intramolecular pairing.

Figure
3.8.
Secondary
structures
of
the
A3
site-containing
region.
The
HIV-1
pre-messenger
RNA
region
containing
the
A3
site
can
adopt
two
different
conformations.
The
A3
site
(shown
in
orange)
containing
the
pyrimidine-rich
sequence
is
accessible
in
structure
1
but
is
not
accessible
in
structure
2
because
it
is
almost
completely
paired
with
another
sequence
(black
nucleotides).
For
a
color
version
of
this
figure,
see

3.3. RNA structures regulating the export of retroviral RNAs
In eukaryotes, almost all cellular pre-messenger RNAs undergo splicing
to remove introns, and only spliced mRNAs can reach the cytoplasm for
translation. Nuclear retention of unspliced or partially spliced transcripts
prevents the synthesis of non-functional or cell-deleterious proteins. In
contrast, the majority of retroviral pre-messenger RNAs must be able to be
exported to the cytoplasm because they have two functions:
– mRNA for the synthesis of Gag and Gag–Pol polypeptide precursors;
– genomic RNA encapsidated in the viral particle.
As a result, retroviruses have evolved strategies to circumvent cellular
restrictions that prevent RNAs containing one or more introns from being
transported from the nucleus to the cytoplasm (Shida 2012). The export
mechanisms of unspliced retroviral RNAs can be divided into two groups
corresponding to complex and simple retroviruses.
3.3.1. Export of unspliced RNA in simple retroviruses
The export mechanisms of unspliced retroviral RNA in simple
retroviruses are different from those used by complex retroviruses. Indeed,
the transport of unspliced RNA from the nucleus to the cytoplasm does not
require a regulatory protein encoded by the retrovirus. Moreover, the NXF1–
NXT1 pathway (also called Tap–p15) is involved in the nuclear export of
unspliced retroviral RNA. This pathway, requiring the formation of a
heterodimer between the NXF1 and NXT1 proteins, is also used for the
transport of cellular mRNAs from the nucleus to the cytoplasm (Katahira
2015). NXF1 is a medium-sized cellular protein (619 amino acids) that has
several functional domains (see Figure 3.9). One, located in the N-terminal
part, corresponding to a nuclear localization signal (NLS), allows NXF1 to be
imported into the nucleus. The N-terminal half of NXF1 contains an RNA
recognition domain (RRM) and a leucine-rich domain (LRR). The binding of
the NXF1–NXT1 heterodimer to RNA involves the RRM, LRR and NTF2L
domains (Katahira 2015). The binding of NXF1–NXT1 to most cellular
mRNAs is not direct; it requires interaction with the TREX1 protein complex
beforehand (Viphakone et al. 2012). The C-terminal half of NXF1, allowing
interaction with the nuclear pore complex (NPC), contains the NTF2L and
UBA domains. NXF1 associates with NXT1 via the NTF2L domain.

Figure 3.9. Structural organization of NXF1. NXF1 has five domains.
The domains are not shown to scale. For a color version of
this figure, see www.iste.co.uk/fosse/structures.zip
Figure 3.10. Unspliced mRNAs of three simple retroviruses. For a color
COMMENTARY ON FIGURE 3.10.– Unspliced mRNAs of three simple
retroviruses belonging to three different genera are shown as examples
(MPMV and SRV-1 for simple betaretroviruses, RSV for alpharetroviruses,
MLV for gammaretroviruses). The cis-regulatory regions of export are in
green and indicated by arrows, except for the one that is present in the U3
region of MLV but has not been precisely localized. The γCTE element is
included in the PTE region of MLV. Regions and genes are not shown to
scale. The cap is indicated by an orange disk.
In simple retroviruses, in contrast to complex retroviruses, several regions
of the unspliced RNA may be involved in its transport (see Figure 3.10).
These cis-regulatory regions may consist of single or multiple stem-loops.
The protein–RNA interactions involved in the export of unspliced RNAs
LRR

from simple retroviruses have not yet been characterized or are only partially
characterized. Nuclear export of unspliced RNAs from MPMV and SRV-1,
two betaretroviruses, is relatively well-understood because it depends on a
single cis-regulatory region (CTE) and the single NXF1–NXT1 pathway.
Nuclear export is more complex in RSV, an alpharetrovirus, because it
depends on two repeat sequences (DR1 and DR2) and, in addition to the
NXF1–NXT1 pathway, the Crm1 pathway could be involved via an
interaction with the viral polypeptide Gag (this point will be discussed in
Chapter 4). The most complex nuclear export is observed in MLV, a
gammaretrovirus, as it involves five regions and also the Crm1 pathway in
addition to the NXF1–NXT1 pathway (Pessel-Vivares et al. 2015; Mougel
et al. 2020).
3.3.1.1. Export of unspliced MPMV and SRV-1 RNAs
Nuclear export of unspliced RNAs from MPMV and SRV-1 retroviruses
depends on the CTE element consisting of 154 nucleotides (Bray et al. 1994;
Zolotukhin et al. 1994). It is likely that the CTE element is of cellular origin
and was captured by a common ancestor of MPMV and SRV-1 (Wang et al.
2015). Specifically, intron 10 of the primate NXF1 gene contains a region of
approximately 100 nucleotides that has high sequence and secondary-
structure homology to the MPMV CTE (Li et al. 2006). This region allows
the nuclear export of NXF1 mRNAs containing intron 10. An in vitro study
using chemical and enzymatic probes showed that the MPMV CTE region
folds into a long stem-loop (Ernst et al. 1997). This stem-loop is
characterized by two inner loops that are arranged in mirror symmetry (see
Figure 3.11).
The use of site-directed mutagenesis has demonstrated that nuclear export
of unspliced RNA is not possible when both inner loops of the CTE of
SRV-1 or MPMV are deleted (Pasquinelli et al. 1997; Grüter et al. 1998).
These studies also showed that the CTE remains functional when it has
only one inner loop. However, the two loops are not equivalent; loop A is
more efficient than loop B for nuclear export. Duplication of the inner
loop in the CTE likely optimizes the export of unspliced RNA (Wang et al.
2015).

Figure 3.11. Secondary structure of the CTE RNA of MPMV. The CTE element forms
a single stem-loop. Arrows indicate identical sequences. The portion above the
dashed line corresponds to the top part of the CTE called hCTE
NXF1 directly binds to each inner loop of the CTE (Grüter
et al. 1998; Kang and Cullen 1999; Teplova et al. 2011). The truncated form
of NXF1, restricted to the RRM and LLR domains (see Figure 3.9), can bind
to the entire CTE or its upper part called hCTE (see Figure 3.11) containing
the inner loop B (Braun et al.1999; Kang and Cullen 1999; Teplova et al.
2011). The three-dimensional structure of the truncated NXF1–hCTE
complex can be determined by X-ray crystallography (Teplova et al. 2011).
The hCTE RNA adopts an L-shaped conformation when it associates with

the truncated NXF1 protein. In this complex, the RRM domain interacts with
the ribose-phosphate backbone constituting the inner loop and the double-
stranded segments that flank it. The LRR domain interacts with bases and
the ribose-phosphate backbone that constitute the single-stranded regions of
the inner loop and the AA bulge. The truncated NXF1–hCTE complex only
partially accounts for the interactions between the whole CTE and the two
NXF1–NXT1 heterodimers. Indeed, the NTF2L domain of NXF1 interacts
with RNA when the protein is associated with NXT1 (Katahira 2015). Thus,
the RRM, LRR and NTF2L domains of NXF1 are involved in CTE element
recognition. To date, there is no three-dimensional structure of the complex
formed by the whole CTE and the two NXF1–NXT1 heterodimers.
Figure 3.12. Secondary structure of a portion of the RSV DR2 RNA. A large portion
of the RSV DR2 element (Prague C strain) folds into a single stem-loop. The
orange-colored guanine plays a crucial role because its replacement by a cytosine
strongly inhibits retrovirus replication and nuclear export. For a color version of this
figure, see www.iste.co.uk/fosse/structures.zip

3.3.1.2. Export of unspliced RNA from RSV
The export of unspliced RNAs from RSV, an alpharetrovirus, is more
complex than that of unspliced RNAs from MPMV. Indeed, RSV unspliced
RNAs appear to use two different pathways to be transported from the
nucleus to the cytoplasm (Maldonado et al. 2020). Although they are
identical, the unspliced RNAs are thought to split into two populations. Each
of the unspliced RNAs in one of the populations functions as a genomic
RNA by associating with the Gag polypeptide, using the Crm1 pathway to
pass into the cytoplasm where it is directed to the plasma membrane and
encapsidated into the virus particle (this process is described in Chapter 4).
The unspliced RNAs from the other population make use of the NXF1 and
Dbp5 (DEAD box RNA helicase) pathway to pass to the cytoplasm, where
they serve as mRNA for Gag or Gag–Pol synthesis (LeBlanc et al. 2007). Both
RSV DR elements (approximately 135 nucleotides each) can be used in export
via the NXF1 pathway, although one is sufficient (Ogert et al. 1996; Simpson
et al. 1997). Since the NXF1 protein does not directly bind to DR1 and DR2
(Paca et al. 2000), it may interact with a previously uncharacterized protein
that binds to DR elements (LeBlanc et al. 2007).
Although the DR1 and DR2 sequences are functionally interchangeable
and show an 82% identity, they do not adopt an identical secondary structure
(Paca et al. 2000). There are, however, a few conserved nucleotides in the 3'
half of the DR elements of different alpharetrovirus strains that are essential
for viral replication and nuclear export (Ogert and Beemon 1998; Yang and
Cullen 1999; Paca et al. 2000; LeBlanc et al. 2007). For example, one of
these corresponds to a guanine in the long stem-loop formed by a large
portion of the DR2 element of the RSV Prague C strain (see Figure 3.12).
3.3.1.3. Export of unspliced RNA from MLV
Although MLV is defined as a simple retrovirus, the nuclear export of its
unspliced RNAs appears to be the most complex in the retrovirus world.
MLV unspliced RNAs use two different pathways for transport from the
nucleus to the cytoplasm (Mougel et al. 2020). The unspliced RNAs are
divided into two populations according to their function in the cytoplasm.
One population corresponds to genomic RNAs that use the Crm1 pathway to
pass into the cytoplasm and are then directed to the plasma membrane where
they are encapsidated in viral particles (Mougel et al. 2020). As in the case
of RSV, the Gag polypeptide could be involved in transport via the Crm1
pathway but this has not been demonstrated. Unspliced RNAs from the other

population make use of the NXF1 pathway to be exported to the cytoplasm
where they serve as mRNA for Gag or Gag–Pol synthesis (Pessel-Vivares et
al. 2014; Sakuma et al. 2014b). The NXF1 protein does not directly bind to
the unspliced mRNA. The nuclear export of MLV unspliced mRNAs via the
NXF1 pathway involves several proteins of the TREX1 complex, which are
the RNA helicase UAP56, THOC5 and THOC7 (Bartels and Luban 2014;
Sakuma et al. 2014a). Unlike UAP56 and THOC7, THOC5 is among the
proteins strongly associated with unspliced mRNA but has not been shown
to directly interact with it. THOC5 may play a role as an adaptor protein by
recruiting UAP56 and THOC7 (Sakuma et al. 2014a).
The molecular mechanisms allowing the nuclear export of the unspliced
RNA of MLV have not been elucidated because they are complex, and it is
difficult to obtain unambiguous results directly linked to an alteration of the
nuclear export with site-directed mutagenesis. Indeed, the unspliced RNA,
corresponding to mRNA or gRNA, is multi-functional. To circumvent these
difficulties, most ex vivo studies have been performed with retroviral vectors
which contain only a part of the viral genome, and thus allow for specifically
studying certain phases of the replicative cycle of a retrovirus. However, the
possibility that an export mechanism characterized with retroviral vectors
does not correspond, or only partially corresponds, to that used by the whole
viral genome cannot be excluded. Ex vivo studies have identified five
cis-regulatory regions of nuclear export of unspliced RNA that are scattered
along the viral genome (see Figure 3.10). The cis-regulatory activity present
in the U3 region has not been shown to be dependent on one or more RNA
structures (Volkova et al. 2014).
The first nuclear export signal was identified using the whole viral
genome and retroviral vectors (Trubetskoy et al. 1999). This signal called
RSL is located at the 5' end of the unspliced RNA (see Figure 3.10) and its
activity is related to its ability to form a stem-loop structure (see
Figure 3.13). The proteins that interact with RSL during nuclear export have
not been characterized.
Two independent studies using retroviral vectors have shown that the
gRNA packaging signal called Psi may also have a role as a cis-regulatory
region for nuclear export of unspliced RNA (Basyuk et al. 2005; Smagulova
et al. 2005). This region folds in the form of four stem-loops (see Figure 3.13).
In the natural context, Psi is probably involved in the nuclear export of
unspliced RNA. Indeed, an overexpression of Psi by a retroviral vector in an

MLV-infected cell competitively leads to a strong decrease in the transport
of unspliced and whole MLV RNA from the nucleus to the cytoplasm
(Smagulova et al. 2005). The proteins involved in this transport have not
been identified. The Gag polypeptide that interacts with Psi during gRNA
encapsidation (a process described in Chapter 4) could be involved and use
the Crm1 pathway but this has not been demonstrated.
A study performed with the whole viral genome and retroviral vectors
identified a long region (1,468 nucleotides) in the MLV pol gene called PTE
(see Figure 3.10) that plays an essential role in the nuclear export of
unspliced RNA (Pilkington et al. 2014). The same study determined the
secondary structure of the PTE region in vitro using two chemical probes.
This folds as seven long stem-loops annotated SL1–SL7 in the 5'–3'
direction. SL1 and SL7 were found to play a major role in the activity of the
PTE region.
Another study, performed with retroviral vectors, characterized the γCTE
element in SL2 (see Figure 3.10), which has similarities to the CTE element
of MPMV (Bartels and Luban 2014). Specifically, the AAGACA sequence
that is present in the apical loop of γCTE is also present in the
NFX1-interacting inner loops of CTE (see Figure 3.13). Although the γCTE
element has not been shown to bind the NXF1 protein, the results indicate
that it contributes to the nuclear export that depends on this protein. The
main function of γCTE is to enable polysome formation during translation of
the gag and gag–pol genes (Bartels and Luban 2014).
A study conducted with retroviral vectors (Sakuma et al. 2014b)
identified another nuclear export cis-regulatory element that is also in the pol
gene, but downstream of the PTE region (see Figure 3.10). This element
called CAE can form three short stem-loops (see Figure 3.13). It facilitates
the nuclear export of unspliced RNA via the NFX1 pathway. A direct
interaction between the NXF1 protein and CAE has not been shown. The
CAE element contains the sequence GGAAAGGAC which is highly
conserved among gammaretroviruses. This sequence plays an essential role
in the CAE activity, but is not sufficient to allow the transport of unspliced
RNA from the nucleus to the cytoplasm.

Figure
3.13.
Secondary
structures
of
nuclear
export
signals
in
MLV.
The
sequence
in
orange
in
the
CAE
element
is
highly
conserved
among
gammaretroviruses.
The
orange
sequence
in
the
γ
CTE
element
is
also
present
in
the
two
inner
loops
of
the
MPMV
CTE.
For
a
color
version
of
this
figure,
see
www.iste.co.uk/fosse/structures.zip.

3.3.2. Unspliced RNA export in complex retroviruses
The mechanisms of unspliced retroviral RNA export are similar in
complex retroviruses belonging to the genera betaretrovirus (JSRV, MMTV,
etc.), deltaretrovirus (HTLV-1, HTLV-2, etc.) and lentivirus (HIV-1, HIV-2,
SIV, etc.). The export of unspliced RNAs, using the Crm1 pathway, depends
on a specific interaction between a regulatory protein encoded by the
retrovirus and an RNA structure formed by a region of the unspliced
retroviral RNA. This cis-regulatory region is located in the 3' part of the
genome, and its length and location vary according to the retroviral species
(see Figure 3.14). In all complex retroviruses, it adopts a conformation
consisting of several stem-loops. The protein–RNA interactions involved in
the export of unspliced RNAs from complex betaretroviruses have not been
characterized, and those occurring in deltaretroviruses are only partially
elucidated. It is in HIV-1 that the export mechanism of unspliced RNA was
discovered (Malim et al. 1989) and has been most extensively studied
(Kuzembayeva et al. 2014).
Figure 3.14. Unspliced mRNAs of three complex retroviruses. For a color
COMMENTARY ON FIGURE 3.14.– Unspliced mRNAs from three complex
retroviruses belonging to three different genera are shown as examples. The
cis-regulatory export regions are shown in green and are called RmRE,
RxRE and RRE in MMTV, HTLV-1 and HIV-1, respectively. RmRE (490
nucleotides) includes the 3' end of the env gene and the 5' end of the U3
region. RxRE (255 nucleotides) includes the 3' end of the U3 region and a
large part of the R sequence. RRE (351 nucleotides) is located in the env
gene. Regions and genes are not shown to scale. The cap is represented by
an orange disk.

3.3.2.1. Export of unspliced HTLV-1 RNA
In deltaretroviruses (HTLV-1, HTLV-2, etc.), it is the Rex protein that
plays an essential role in the export of unspliced or partially spliced viral
RNAs (Nakano and Watanabe 2016). Like the Rev protein of lentiviruses,
Rex is a regulatory protein that is expressed during the early phase of viral
replication. It is translated from doubly spliced viral mRNAs that, like those
encoding the Tax protein (equivalent to the HIV-1 Tat protein), are the first
to be transported to the cytoplasm after transcription of proviral DNA. These
spliced viral mRNAs are transported from the nucleus to the cytoplasm by
the same pathway used by cellular mRNAs.
Figure 3.15. Structural organization of Rex. The N-terminus contains the arginine-
rich domain that includes the NLS. The bipartite oligomerization domain and the
nuclear export signal (NES) are in the central portion. The C-terminus contains the
domain that stabilizes the protein. The domains are not shown to scale. For a color
Rex is a small protein (189 amino acids in HTLV-1) that has several
functional domains (see Figure 3.15). One, located in the N-terminal part,
allows Rex to be imported into the nucleus and to bind to RNA because it
contains an NLS and is rich in arginines (D’Agostino et al. 2019). The
central part of Rex contains a bipartite oligomerization domain and an NES.
Through NLS and NES signals, Rex can shuttle between the cytoplasm and
nucleus (Nakano and Watanabe 2016). The C-terminus corresponds to a
domain that stabilizes the protein by increasing its half-life (Xie et al. 2009).
The export of unspliced and partially spliced viral RNAs requires Rex
binding to the RxRE region (255 nucleotides) (Nakano and Watanabe 2016).
Note that, unlike the HIV-1 cis-regulatory RRE sequence, RxRE is present
in all HTLV-1 RNAs because it is located downstream of splice acceptor
sites and in the 3' end of the genome (see Figure 3.14). RNAs whose export
depends on their interaction with Rex have a 3' CRS element that is
responsible for their retention in the nucleus and degradation in the absence
of Rex (Cavallari et al. 2016). By binding to RxRE, Rex abolishes the
negative effect of CRS by enabling transport from the nucleus to the
cytoplasm via the Crm1 pathway. The RxRE RNA forms a secondary
structure with four stem-loops (see Figure 3.16) (Toyoshima et al. 1990;
Askjaer and Kjems 1998). Of these four stem-loops, stem-loop D is
ARG/NLS OLIGO NES OLIGO STAB

primarily responsible for the nuclear export activity of RxRE (Toyoshima
et al. 1990; Gröne et al. 1994). However, the other three stem-loops must be
present so that RxRE RNA folds into the form of the four stem-loop
structure, which is optimal for nuclear export. One hypothesis is that this
structure stabilizes stem-loop D and/or interacts with unidentified nuclear
factors that facilitate Rex-dependent mRNA transport.
Figure 3.16. RxRE RNA folding. Diagram of RxRE folding
consisting of stem-loops A–D and stems I and II
The formation of the Rex–RxRE–Crm1 complex has been much less
studied than the equivalent HIV-1 complex. Therefore, there is no model
describing in detail the steps leading to the formation of the Rex–RxRE–
Crm1 complex. However, studies of the Rex–RxRE interaction have shown
that Rex binds only to stem-loop D (Bogerd et al. 1991; Askjaer and Kjems
1998). Rex protein can bind to the isolated stem-loop D but its affinity for it
is greatly diminished compared to the affinity it has for the entire RxRE

region (Askjaer and Kjems 1998). The four-loop stem folding of RxRE thus
plays an important role in its interaction with Rex. Two studies (Bogerd
et al. 1991; Baskerville et al. 1995) have identified a two-nucleotide bulge
(see Figure 3.17, site 1) as the unique site of interaction with Rex. A second
high-affinity site (site 2) was characterized by another study that was
performed under different experimental conditions than the previous two
studies (Askjaer and Kjems 1998). Stem-loop D therefore binds at least two
Rex proteins and no more than four through protein–protein interactions
a priori. Crm1 could induce Rex oligomerization (Hakata et al. 1998). To
date, the process of Crm1–Rex–RxRE complex formation has not been
determined.
Figure 3.17. Secondary structure of stem-loop D. This diagram shows the stem-loop
D of the RxRE region that plays a crucial role in Rex binding. The two sites with high
affinity for Rex are circled in pink. Site 1 has been identified in several studies, while
site 2 has been characterized in only one study. For a color version of this figure, see
3.3.2.2. Export of unspliced HIV-1 RNA
The Rev protein of HIV-1 plays a crucial role in the export of unspliced
or partially spliced RNAs from this virus (Malim et al. 1989; Kuzembayeva
et al. 2014). Rev is a regulatory protein that is expressed during the early
phase of viral replication. Indeed, it is translated from spliced viral mRNAs
of about 2 kb that, like those encoding the Tat protein, are the first to be
transported to the cytoplasm after transcription of the proviral DNA. These
spliced viral mRNAs use the same export pathway to the cytoplasm as that
used by cellular mRNAs.
Figure 3.18. Structural organization of Rev. The half of the protein that is structured
contains the bipartite oligomerization domain and the arginine-rich domain that
includes the NLS. The other half that is disordered contains the NES. The domains
are not shown to scale. For a color version of this figure, see www.iste.co.uk/fosse/
structures.zip
OLIGO ARG/NLS OLIGO NES

Rev is a small protein (116 amino acids) whose N-terminal half is
structured, while the C-terminal half is disordered. This protein, despite its
small size, has several functional domains (see Figure 3.18). Indeed, Rev
contains an NLS that allows it to be imported into the nucleus. This signal is
included in an arginine-rich domain, enabling Rev to bind to RNA. Rev
also contains a bipartite oligomerization domain and a leucine-rich NES.
Through the NLS and NES signals, Rev can shuttle between the cytoplasm
and nucleus (Rausch and Le Grice 2015).
Transport from the nucleus to the cytoplasm of unspliced and partially
spliced viral RNAs (approximately 4 kb in size) requires the interaction of
Rev with the RRE region (351 nucleotides) that is present in these RNAs
(Malim et al. 1989; Rausch and Le Grice 2015). RRE can form two
secondary structures that are approximately in equimolar amounts (Watts
et al. 2009; Rausch and Le Grice 2015; Sherpa et al. 2015). In both
structures, the 3' and 5' ends form a long stem that extends into a four or five
stem-loop conformation (see Figure 3.19). The two structures differ only in
stem-loops III and IV. The five stem-loop conformation is the one required
for optimal HIV-1 replication (Sherpa et al. 2015). The RRE regions of SIV
and HIV-2 lentivirus also adopt five stem-loop conformations (Lusvarghi
et al. 2013; Pollom et al. 2013). The RRE region of HIV-1 has been studied
twice using the small-angle X-ray scattering (SAXS) technique, which
provides information on the overall shape of the RNA. One of the studies
was performed with the truncated RRE region at the lower part of stem I
(Fang et al. 2013), while the other was performed with the entire RRE region
(Bai et al. 2014). The truncated RRE region adopts a three-dimensional
folding that is A-shaped, while the three-dimensional folding of the whole
RRE region resembles a modified A (see Figure 3.20). These conformations
result from coaxial stacking between the stems of the RRE region.
The binding of Rev to RRE is cooperative and sequential. Rev
oligomerizes onto RRE one molecule at a time. In a first step, one molecule
of Rev binds first to the site in stem IIB and then forms a dimer with the
second Rev molecule which binds at the junction of the IA, IIB and IIC
stems. Oligomerization of Rev continues to form an RRE–Rev complex
consisting of one RRE molecule and at least six Rev molecules (Daugherty
et al. 2010a). The determination of the exact number of Rev by RRE is a
scientific controversy, as the result obtained differs depending on the
experimental methods used (Rausch and Le Grice 2015).

Figure 3.19. Secondary structures of the RRE. For a color version
COMMENTARY ON FIGURE 3.19.– This diagram shows the part of the RRE
region that plays a crucial role in the attachment of Rev. Stem I of the entire
RRE region is much longer than that shown here. RRE can form a secondary
structure with five stem-loops (left) or four stem-loops (right). The base
pairing interactions between stems IIA and V are different in the two
conformations. The three sites with high affinity for Rev are circled in pink.
Figure 3.20. Models of the three-dimensional folding of RRE
III-V
IIB
IIC
I
IA
III-V
IIB
IIC
I
I

COMMENTARY ON FIGURE 3.20.– The three-dimensional foldings of the
truncated and whole RRE regions are schematized by an A (left) and a
modified A (right), respectively. In the whole RRE region, the long stem I,
which is much shorter in the truncated RRE region, folds and establishes
tertiary interactions with nucleotides forming the stem IA. The stems (I, IA,
etc.) constituting the topological elements are indicated within them.
A model for the formation of the Rev–RRE complex (see Figure 3.21)
was proposed based on SAXS data and analysis of Rev binding to the whole
RRE RNA using the SHAPE method (Bai et al. 2014). The model is
characterized by a change in the three-dimensional folding of RRE induced
by the formation of the Rev tetramer. This conformational change makes a
site of Rev accessible that is located in stem I.
Figure 3.21. Model of Rev binding to preorganized RRE RNA. The RRE RNA,
having a preorganized three-dimensional A-shaped structure, guides the sequential
binding of Rev until a hexamer is formed that is competent to bind a Crm1–Ran
GTP
dimer via two of its six NESs. Each Rev molecule is represented by a yellow oval
shape. For a color version of this figure, see www.iste.co.uk/fosse/structures.zip
To date, the three-dimensional structure of the large and flexible
Rev–RRE–Crm1–RanGTP
complex has not been determined. However,
a structural model of the Rev–RRE–Crm1–RanGTP
complex has been
proposed based on biochemical data and electron microscopy data and
through the determination by X-ray crystallography of the structure of the
dimer formed by Rev (Daugherty et al. 2010b; Booth et al. 2014).
In this jellyfish-like model, one side of the Rev hexamer interacts with
RNA via the arginine-rich region, while the opposite side projects outward
the six tentacles consisting of the six disordered C-termini, each containing

an NES (see Figure 3.22). The Rev–RRE complex binds a Crm1–RanGTP
dimer via two of its six NESs and can thus pass through the nuclear pore.
Figure 3.22. Model of the Rev–RRE–Crm1–Ran
GTP
complex. For a color
COMMENTARY ON FIGURE 3.22.– The hexamer of Rev adopting a jellyfish-
like structure is colored yellow. The structured part of each subunit of the
hexamer is schematized by a cylinder, while the disordered tentacle-like part
corresponds to a thick line. The secondary structure of RRE RNA forming
the five stem-loop conformation is in blue. Each subunit of the Crm1 dimer
is represented by a gray oval shape. The RanGTP
associated with Crm1
corresponds to the black oval shape. The two purple disks indicate the two
sites through which the Rev–RRE complex is tied to the Crm1–RanGTP
dimer.
3.4. RNA structures regulating the translation of retroviral RNAs
In eukaryotic cells, translation is a process that can be divided into four
main phases: initiation, elongation, termination and recycling of ribosomes.
Translational regulation is mainly carried out at initiation and consists of
several steps leading to the assembly of the 80S ribosome at the translation

start codon (almost exclusively the AUG triplet). In the vast majority of
cases, the initiation of translation is dependent on the 5' cap (mGppp7
). This
recruits the 40S subunit of the ribosome via its interaction with the
translation initiation factor eIF4E. Cap-independent translation initiation has
been discovered in picornaviruses, whose mRNAs do not have a cap at their
5' end (Jang et al. 1988; Pelletier and Sonenberg 1988). In picornaviruses,
translation initiation depends primarily on a structured region of the mRNA
that constitutes an internal ribosome entry site (IRES). Ten to fifteen percent
of cellular mRNAs contain an IRES, although they possess a cap (Barrera
et al. 2020a). The presence of a cap and an IRES on the same mRNA allows
for a switch from cap-dependent to IRES-dependent translation initiation
when the cell is under stress. The activity of an IRES requires it to interact
with factors called IRES trans-acting factors (ITAFs). ITAFs are usually
chaperone proteins that allow an IRES to adopt the functional
conformation.
Retroviruses, all of whose mRNAs possess a cap, use the translation
machinery of the eukaryotic cells they infect. IRESs have been identified in
both simple and complex retroviruses (Barrera et al. 2020a). Translation
initiation of retroviral mRNAs has been studied mainly in HIV-1 and HIV-2,
which are complex viruses. However, a few studies on translation initiation
have been performed with simple retroviruses.
3.4.1. IRESs of simple retroviruses
The first retroviral IRESs were identified in simple retroviruses belonging
to the genera gammaretrovirus and alpharetrovirus (Berlioz and Darlix 1995;
Vagner et al. 1995; López-Lastra et al. 1997; Deffaud and Darlix 2000a,
2000b). They are in the structured regions that are formed by the 5'-UTR
domains of retroviral mRNAs. With the exception of the IRES that is present
in the F-MLV env mRNA, the IRESs which have been characterized are
located in the unspliced mRNAs (see Figure 3.23). A common structure for
the different IRESs was not identified in the 5'-UTR regions of the simple
retroviruses studied.

Figure 3.23. Localization of IRES in single retrovirus mRNAs. For a color
COMMENTARY ON FIGURE 3.23.– The gag–pol mRNAs of gammaretroviruses
(MoMuLV and REV-A) and an alpharetrovirus (RSV) are shown as
examples. The env mRNA of F-MLV, a gammaretrovirus, is also shown.
IRESs, green regions, are indicated by arrows. Regions and genes are not
shown to scale. The cap is indicated by an orange disk.
3.4.1.1. IRESs of gammetrovirus
The 5'-UTR domain of the unspliced mRNAs of two murine retroviruses
(MoMuLV and F-MLV) and one avian retrovirus (REV-A) has IRES
activity (Berlioz and Darlix 1995; Vagner et al. 1995; López-Lastra et al.
1997). This activity depends mainly on a region just upstream of the gag
gene initiation codon, which consists of 126 (MoMuLV) or 129 (REV-A)
nucleotides (Vagner et al. 1995; López-Lastra et al. 1997). In the case of
MoMuLV, part of this region can form two stem-loops, one of which carries
a pyrimidine motif (see Figure 3.24) that plays an important role in IRES
activity. An interaction between this motif and the splicing factor PTB has
been demonstrated (Vagner et al. 1995). Interestingly, the IRES activity of
several picornaviruses requires a pyrimidine and PTB motif that is part of
the ITAFs (Barrera et al. 2020a). Other ITAFs involved in the IRES activity
of unspliced gammaretrovirus mRNAs have not been identified. The study
that identified an IRES in the 5'-UTR domain of F-MLV env mRNA is the
only one to date that shows that the initiation of translation of a
gammaretrovirus mRNA can depend on an IRES (Deffaud and Darlix
2000a). This IRES, whose secondary structure has not been determined, is

located between the splice donor/acceptor site and the initiation codon of the
env gene (see Figure 3.23). The possibility that the mRNAs of many simple
retroviruses possess an IRES cannot be ruled out, because the IRES activity
of these mRNAs has hardly been investigated.
Figure 3.24. Secondary structure of the 3' end of the IRES of the MoMuLV unspliced
mRNA. The 3' end of the IRES of MoMuLV unspliced mRNA can form two
stem-loops (Mougel et al. 1993). The pyrimidine motif interacting with PTB is in
orange. The initiation codon of the gag and gag–pol genes is in black. For a color
3.4.1.2. IRESs of an alpharetrovirus
The 5'-UTR region of the unspliced mRNA of RSV, an alpharetrovirus,
contains an IRES that is bipartite (see Figure 3.23). Both parts, when
isolated, each exhibit IRES activity, which is, however, weaker for the 5'
part than for the 3' part (Deffaud and Darlix 2000b). The secondary
structures of the 5'-UTR region and the ITAFs that are required for IRES
activity have not been characterized.
3.4.2. Translation initiation in complex retroviruses
Apart from lentiviruses, and more particularly HIV-1, few studies have
been devoted to the identification of mechanisms regulating the initiation of
translation of complex retroviruses mRNAs. With the exception of HIV-2,
all of the complex retroviruses studied have an IRES in the 5'-UTR domain
of the unspliced mRNA (see Figure 3.25).

Figure 3.25. Localization of IRESs in unspliced mRNAs of complex retroviruses.
For a color version of this figure, see www.iste.co.uk/fosse/structures.zip
COMMENTARY ON FIGURE 3.25.– The gag–pol mRNAs of MMTV, HTLV-1,
HIV-1 and HIV-2 retroviruses are shown as examples. IRESs in the 5'-UTR
domains are in green. IRESs in the gag gene are in orange. In the case of
HIV-2, the orange part corresponds to three IRESs. Regions and genes are
not shown to scale. The cap is indicated by an orange disk.
3.4.2.1. IRESs of MMTV
The 5'-UTR domain (320 nucleotides) of the unspliced mRNA of MMTV
has an IRES (Vallejos et al. 2010). It has not been precisely localized, and its
secondary structure has not been determined. PTB, which binds to the
5'-UTR region and stimulates IRES activity, has been identified as an ITAF
(Cáceres et al. 2016). The hnRNP A1 protein has been characterized as
another ITAF, as it is required for IRES activity (Barrera et al. 2020b).
3.4.2.2. IREs of HTLV-1
The 5'-UTR domain of the HTLV-1 unspliced mRNA contains a
181-nucleotide long IRES that is just upstream of the gag gene (Olivares et
al. 2014). Its secondary structure has not been determined. IRES activity is
dependent on the ribosomal protein S25 and is stimulated by the hnRNP A1
protein, which is an ITAF (Olivares et al. 2014; Barrera et al. 2020b).
The HBZ regulatory protein is encoded by the negative polarity strand of
the HTLV-1 genome. Two isoforms of HBZ are synthesized from two
antisense mRNAs, which are either spliced or unspliced. The first 218

nucleotides of the 5'-UTR region of the spliced antisense mRNA constitute
an IRES (Cáceres et al. 2018). The IRES folds as three domains (see
Figure 3.26). Domain I plays the most important role in IRES activity. This
is dependent on the ribosomal protein S25 and the translation initiation
factor eIF5A. Interactions of these or other proteins with IRES have not been
reported in the studies published to date.
Figure 3.26. Folding of the 5'-UTR domain of spliced HBZ mRNA. The IRES, shown
in green, consists of domains I, II and III. Domain I contains two stem-loops and a
large inner loop. Domain IV, in blue, corresponds to the 3' end of the 5'-UTR that is
just upstream of the AUG codon of the HBZ gene. For a color version of this figure,
3.4.2.3. IRESs of HIV-2
To date, there are no studies that have focused on the initiation of
translation of HIV-2-spliced mRNAs. HIV-2 differs in its mode of initiation
of unspliced mRNA translation from other simple and complex retroviruses
that have been studied. Indeed, IRES activity in the 5'-UTR domain of the
HIV-2 unspliced mRNA could not be demonstrated (Barrera et al. 2020a).

Furthermore, translation initiation does not appear to be dependent on the
presence of the 5'-cap at any point in the replicative cycle of the virus.
The initiation of translation of the unspliced HIV-2 mRNA depends
primarily on the three IRESs that are located in the 5' end of the coding
portion of the gag gene (Herbreteau et al. 2005; Ricci et al. 2008; Weill et al.
2010). The region containing these IRESs is approximately 350 nucleotides
long and folds into six main domains (see Figure 3.27). The first IRES is
downstream of the AUG1 initiation codon where synthesis of the Gag and
Gag–Pol polypeptides begins. The initiation of translation of these
polypeptides is original, as it depends on an IRES that is downstream and
not upstream of the AUG codon. The AUG2 and AUG3 initiation codons
allow the synthesis of truncated Gag polypeptides whose roles in virus
replication are not known. The second IRES is located between AUG1 and
AUG2. The third IRES is between AUG2 and AUG3 (Ricci et al. 2008).
The three IRESs function independently and are able to recruit three
ribosomal translation initiation complexes to the same unspliced mRNA
molecule (Weill et al. 2010). The region containing domains P2–P5 (see
Figure 3.27) is capable of directly binding the 40S subunit of the ribosome
and the translation initiation factor eIF3 (Locker et al. 2011). The interaction
sites have not been precisely identified. These results led to the proposal of a
two-step sequential process that occurs three times to allow three initiations
on the same RNA molecule:
– recruitment by the P2–P5 region of the 40S subunit and the eIF3 factor;
– transfer of the 40S subunit and the eIF3 factor to one of the three AUG
codons.
The molecular basis of the mechanism involved in the transfer has not
been determined. The transfer may require the disruption of intramolecular
base pairing interactions within IRESs because the synthesis of whole and
truncated Gag polypeptides requires the presence of eIF4A, which is an
RNA helicase (Locker et al. 2011).

Figure 3.27. Folding of the 5' end of the HIV-2 gag gene. For a color
COMMENTARY ON FIGURE 3.27.– The synthesis of Gag and Gag–Pol
polypeptides starts at codon AUG1. The synthesis of truncated Gag
polypeptides is initiated at codons AUG2 and AUG3. The three IRESs are in
the highly structured region outlined in green. The 5'-UTR region is
upstream of the AUG1 codon. The sequence downstream of codon AUG3
corresponds to the 3' part of the gag gene.
3.4.2.4. IRESs of HIV-1
Three different mechanisms may be responsible for the initiation of
translation of unspliced HIV-1 mRNA:
A
U
G 1
U
A
C
.
.
.
.
.
.
. . .
5’
P1
P2
P3
A
U
G 2
P4
P5
. . . 3’
A
U
G3
P6

– canonical initiation dependent on the 5' cap and its direct interaction
with eIF4E;
– non-canonical initiation dependent on the 5' cap but not requiring
interaction of the latter with the eIF4E factor;
– non-canonical initiation via IRESs (Barrera et al. 2020a).
Studies have shown that the initiation of translation is a dynamic process
that is primarily cap-dependent at the onset of HIV-1 infection of the cell
(Gendron et al. 2011; Vallejos et al. 2011; Monette et al. 2013). Over time,
infection induces a cellular environment (e.g. oxidative stress) that is
unfavorable for cap-dependent initiation but allows initiation via IRES.
The TAR stem-loop, which is located at the 5' end of unspliced and
spliced mRNAs, is conserved in different HIV-1 isolates because it plays a
crucial role in proviral DNA transcription (see section 3.1.2). However, it
inhibits translation in vitro because, due to its high stability, it is a barrier
during cap recruitment of eIF4E and other factors forming the translation
preinitiation complex with the 40S subunit of the ribosome (de Breyne and
Ohlmann 2018). The virus can use ex vivo cellular proteins (Staufen-1,
UPF1, TRBP, etc.) and viral proteins (Rev and Tat) to overcome the
inhibition induced by the TAR structure. These proteins’ modes of action
have not all been characterized. However, it is known that several of these
proteins bind to the TAR and that some possess RNA helicase activity
(de Breyne and Ohlmann 2018; Barrera et al. 2020a). They thus act by
destabilizing the TAR structure and therefore allowing the translation
preinitiation complex to form at the cap and slide along the mRNA.
The initiation of unspliced mRNA translation becomes cap-independent
and dependent on two IRESs, which are located in the 5'-UTR domain and
the gag gene, respectively (see Figure 3.25), when the cellular environment
changes and the G2/M transition of the cell cycle is blocked by the HIV-1
Vpr protein (Brasey et al. 2003; Barrera et al. 2020a). The 5'-IRES
(approximately 225 nucleotides), which is present in the 5'-UTR region and
just upstream of the gag gene initiation codon, enables the synthesis of Gag
and Gag–Pol polypeptides (Brasey et al. 2003; Gendron et al. 2011; Vallejos
et al. 2011). It consists of the four structured domains PBS, DIS, SD and Psi
(see Figure 3.28). It is probably the topological arrangement of the four
domains that confers IRES activity (Gendron et al. 2011; Vallejos et al.

2011; Plank et al. 2013). The 5'-IRES requires cellular proteins to be
functional. Among these, the hnRNP A1 protein was the first ITAF to be
characterized (Monette et al. 2009). This protein’s mechanism of action,
which stimulates the 5'-IRES activity, has not been determined. The cellular
protein HuR is an ITAF that inhibits the 5'-IRES activity (Rivas-Aravena
et al. 2009). Other cellular proteins (DDX3 helicase and hRIP) have been
characterized as ITAFs (Barrera et al. 2020a). The molecular mechanisms
that are used by ITAFs to regulate the 5'-IRES activity are not known. The
ribosomal protein S25 is involved in the recruitment of the translation
preinitiation complex by the 5'-IRES (Carvajal et al. 2016). A direct
interaction between this protein and the 5'-IRES probably does not occur as
it could not be demonstrated.
Figure 3.28. Folding of the 5'-UTR domain of HIV-1 unspliced mRNA. For a
COMMENTARY ON FIGURE 3.28 – The 5'-UTR region of the unspliced mRNA
can fold into six stem-loops (Wilkinson et al. 2008) of which the two closest
to the 5' end are not required for IRES activity. The IRES in green is
upstream of the gag gene initiation codon (AUG in black). The apical loop
of SL2 contains the splice donor site (SD).
5’
TAR
poly(A)
A
U
G
PBS
SL3
.
.
.
3’
SL1
SL2

AUG
5’…
336
G
U
A
759
3’
Site
1
Site
2
Figure
3.29.
HIV-1
IRES–Gag
folding.
For
a
color
version
of
this
figure,
see

COMMENTARY ON FIGURE 3.29.– The IRES–Gag is located between codons
AUG 336 and 759. Codon AUG 336 allows the synthesis of the Gag
polypeptide, while codon AUG 759 allows the synthesis of the Gag–p40
protein. Sites 1 and 2 shown in orange correspond to the two sites where the
40S subunit of the ribosome binds. These sites contain an adenine-rich
region, which is boxed in pink.
The IRES–Gag (approximately 420 nucleotides), which is located in the
gag gene (see Figure 3.25), synthesizes the Gag polypeptide and a truncated
form of the Gag polypeptide, which is called Gag–p40 (Buck et al. 2001).
The role of the Gag–p40 protein is not known, but it is known that virus
infectivity decreases if the protein is not produced. As in HIV-2, the highly
structured IRES–Gag (see Figure 3.29) is located downstream of the gag
gene initiation codon. Two sites binding the 40S subunit of the ribosome
have been identified in the IRES–Gag (Deforges et al. 2017). Each site is
thought to recruit a 40S subunit of the ribosome via an adenine-rich
sequence. Gag–p40 synthesis results from preinitiation complex sliding from
site 2 to AUG 759 in the canonical 5'–3' direction. In contrast, a preinitiation
complex shift occurs from site 1 to AUG 336 in the non-canonical 3'–5'
direction to enable the synthesis of the Gag polypeptide (Deforges et al.
2017). ITAFs involved in IRES–Gag activity have not been identified. To
date, we do not know how the 5'–IRES and IRES–Gag coordinately regulate
Gag polypeptide synthesis from the same AUG codon.
Alternative splicing in HIV-1 produces the nef, tat, vif, vpr and vpu
mRNAs, which also have an IRES in their 5'-UTR domain (Charnay et al.
2009; Plank et al. 2014). All of these mRNAs share in common with the
unspliced mRNA the same sequence (289 nucleotides), which is upstream of
the splice donor site (SD) and constitutes part of the 5'-UTR domain. The
folding of this sequence, which is the same in the different mRNAs, leads to
the formation of the TAR, poly(A), PBS and DIS (Plank et al. 2014). IRES
activity does not require the TAR and poly(A) stem-loops but does require
the PBS and DIS structures. IRES activity is modulated by the sequence that
is downstream of the splice acceptor site and corresponds to the other part of
the 5'-UTR domain. ITAFs that are associated with the 5'-IRES of mRNAs
have not been characterized.
The 5' part of exon 1 of tat has a short sequence (16 nucleotides), whose
stem-loop structure called TIM-TAM is highly conserved among different
HIV-1 isolates (Khoury et al. 2020). This stem-loop corresponds to an

independent structural domain in the gRNA (see Figure 3.30), while it
constitutes the apical part of a long stem-loop in the tat mRNA. It plays a
dual role (Khoury et al. 2020):
– it is necessary for the IRES activity which is present in the 5' coding
end of the tat gene;
– it blocks the progression of the ribosome when the initiation of the
translation of tat mRNA depends on the cap.
The proteins involved in IRES activity have not been identified. It has
been proposed that in the early phase of infection prior to viral DNA
integration, initial Tat protein synthesis is dependent on TIM-TAM, which is
present in the gRNA (Khoury et al. 2020).
Figure 3.30. Secondary structure of TIM-TAM in gRNA.
The stem-loop TIM-TAM is flanked by two short, unpaired
sequences in the gRNA (Watts et al. 2009)

4
Encapsidation of Genomic
RNA in the Retroviral Particle
Studies performed with alpharetroviruses, gammaretroviruses,
lentiviruses and various techniques (analytical ultracentrifugation, electron
microscopy and agarose gel electrophoresis) showed that the gRNA
extracted from the virions is in the form of a dimer whose subunits are
linked by non-covalent bonds (Mangel et al. 1974; Bender and Davidson
1976; Bender et al. 1978; Murti et al. 1981; Fu et al. 1994; Höglund et al.
1997). The virion of all retroviral species thus contains a diploid genome that
consists of two gRNA molecules. The reverse transcription of the diploid
genome generates genetic diversity that allows retroviruses to develop
resistance to antiretrovirals and to evade the host immune system (see
Chapter 2, section 2.4). With the exception of spumaretroviruses,
encapsidation is a process common to all retroviruses that selects and
packages into a viral particle two gRNA molecules from the majority of viral
and cellular mRNAs. This process requires the binding of the Gag precursor,
via its NC domain, to the gRNA. In several retroviral species, the gRNA
must first dimerize in order to be recognized by the Gag polypeptide.
4.1. RNA structures and mechanisms governing gRNA
dimerization
The gRNA dimer extracted from virions (mature virus particles) is
described as stable because dissociation of the subunits requires a relatively
high temperature. In contrast, the gRNA dimer extracted from non-infectious
immature viral particles is considered unstable because dissociation of the

subunits occurs at a significantly lower temperature (Stoltzfus and Snyder
1975; Fu and Rein 1993; Fu et al. 1994). Conversion of the unstable dimer
to the stable dimer requires cleavage of the Gag polypeptide by the viral
protease (Stewart et al. 1990; Fu and Rein 1993; Fu et al. 1994;
Shehu-Xhilaga et al. 2001b). Studies based on site-directed mutagenesis
suggest that NC, a cleavage product of the Gag precursor, plays an important
role in stabilizing the gRNA dimer (Méric and Spahr 1986; Méric et al.
1988; Fu and Rein 1993).
Electron microscopy has enabled the identification of the main interaction
between the subunits of the gRNA dimer that is extracted from the virion
(Bender and Davidson 1976; Murti et al. 1981; Höglund et al. 1997). This
interaction, termed DLS, occurs near the 5' end of the gRNA in the region
containing the encapsidation signal. Since the electron microscopy studies
were performed under semi-denaturing conditions, we cannot completely
exclude the possibility that these conditions induce conformational changes
in the gRNA, allowing it to form a DLS that does not exist in the virion.
Dimerization of gRNA from several retroviruses has been reproduced in
vitro using short RNAs that were synthesized in vitro and corresponded to
the 5' ends of retroviral genomes (Bieth et al. 1990; Darlix et al. 1990; Prats
et al. 1990; Roy et al. 1990; Marquet et al. 1991). Numerous in vitro studies
have characterized the sequences, structures and mechanisms that regulate
gRNA dimerization in several retroviruses.
4.1.1. Structures and mechanisms in alpharetroviruses
The gRNA dimer of RSV type Pr-A, an alpharetrovirus, was extracted
from the virion and analyzed with electron microscopy (Murti et al. 1981).
This analysis indicates that the DLS that associates the two subunits is
located approximately 479–539 nucleotides from the 5' end (see Figure 4.1).
Dimerization of alpharetrovirus gRNA was studied in vitro using RNAs
containing the DLS. In vitro experiments performed under physiological
conditions (temperature and salinity), and with RNAs corresponding exactly
to the 5' part of alpharetrovirus (ALV and RSV) gRNA, could not reveal a
dimerization site at the DLS (Polge et al. 2000; Liu et al. 2020). In contrast,
a contact point associating the two subunits of the RNA dimer generated
in vitro was identified in the L3 element of ALV (SR-A type) that is

Encapsidation of Genomic RNA in the Retroviral Particle 75
approximately 200 nucleotides upstream of the DLS (Fossé et al. 1996;
Polge et al. 2000). The L3 element adopts a stem-loop structure (see
Figure 4.2) that is conserved among alpharetroviruses. A single-base
mutation in the apical loop autocomplementary sequence is sufficient to
suppress dimerization in vitro (Polge et al. 2000). Deletion of the L3 element
decreases replication of SR-A RSV by 20- to 300-fold (Doria-Rose and
Vogt 1998). In addition, there is selection pressure to restore an
autocomplementary sequence in the apical loop after several replication
cycles when it has been mutated (Doria-Rose and Vogt 1998). The
importance of the L3 element in the replication of SR-A-type
alpharetroviruses is probably due to its role in gRNA dimerization.
However, the very low level of replication in the absence of the L3 element
suggests the existence of one or more secondary dimerization sites. In vitro
dimerization of an RNA corresponding to the 5' part of the Pr-C RSV
genome is not solely dependent on the L3 element. Indeed, the SLA and L3
elements must be simultaneously deleted to suppress in vitro dimerization
(Liu et al. 2020). Although the SLA element also corresponds to a stem-loop
whose apical loop contains a self-complementing sequence, it has not been
shown to be a contact point between the two subunits of the RNA dimer.
Furthermore, the infectivity of RSV (Pr-C type) is not diminished by two
deletions that delete the SLA and L3 elements (Liu et al. 2020). Thus, it
appears that the L3 stem-loop is not equally important in different
alpharetrovirus types.
Figure 4.1. Sites involved in RSV gRNA dimerization. For a color
COMMENTARY ON FIGURE 4.1.– The DLS identified by electron microscopy
is in the gag gene downstream of the splice donor site (SD). The SLA and L3
elements, which are upstream of SD, were identified in vitro using an in vitro
synthesized RNA that corresponds to the 5' end of the RSV gRNA. The
orange disk represents the 5' cap. Regions and genes are not shown to scale.

Figure 4.2. Dimerization of alpharetrovirus gRNA via the L3 element. The secondary
structure of the L3 stem-loop in the gRNA monomer is shown in the upper part of the
figure. Shown in blue and black are the two L3 stem-loops that link the stable and
unstable dimer subunits via Watson–Crick base pairs. For a color version of this
Under physiological temperature and salinity conditions, and in the
absence of NC, an RNA, corresponding to the 5' part of ALV gRNA (SR-A
type), forms an unstable dimer in vitro via the L3 element (Polge et al.
2000). This dimer may correspond in part to the gRNA dimer present in the
immature virus particle. The subunits of this dimer dissociate when analyzed
by agarose gel electrophoresis at room temperature and in the absence of
magnesium. The apical loop of the L3 stem-loop is directly involved in the
loop–loop interaction that links the subunits of the unstable dimer (Polge
et al. 2000). This loop–loop interaction, forming a short duplex of six base
pairs (see Figure 4.2), is thought to initiate the formation of a long duplex
linking the subunits of the stable dimer that is present in the mature virus
particle. It is possible to generate a stable dimer using an elevated
temperature (60°C) or physiological temperature in the presence of NC
(Polge et al. 2000; Ben Ali et al. 2007). The subunits of the stable dimer are
linked by a duplex of 16 base pairs that results from the chaperone activity

of the NC. Indeed, it opens the apical part of the L3 stem-loop and promotes
the formation of a long stable duplex that contains more paired nucleotides
than the same sequence in the monomer (see Figure 4.2). Although this
duplex could also be generated by the NC in the virion, this has so far
neither been demonstrated nor refuted.
4.1.2. Structures and mechanisms in betaretroviruses
4.1.2.1. Dimerization of MPMV gRNA
The gRNA dimer of MPMV, which is present in the virion, has not been
studied by electron microscopy. By analogy with other retroviral species, it
is assumed that the 5' part of the genome is responsible for its dimerization.
In favor of this hypothesis, an in vitro synthesized RNA corresponding to the
5' part of the MPMV genome can form a dimer in vitro (Aktar et al. 2013).
The Pal SL element was identified as the contact point associating the two
subunits of the in vitro generated RNA dimer (see Figure 4.3). Although a
35-nucleotide deletion encompassing the Pal SL element decreases MPMV
replication by ninefold, this has not been shown to be due to altered gRNA
dimerization (Jaballah et al. 2010).
Figure 4.3. Site involved in the in vitro dimerization of MPMV gRNA. The Pal SL
element upstream of the splice donor site (SD) was identified in vitro using an in vitro
synthesized RNA that corresponds to the 5' end of MPMV gRNA. The orange disk
represents the 5' cap. Regions and genes are not shown to scale. For a color version
The Pal SL element forms a short stem-loop that has a self-
complementary sequence in the apical loop (see Figure 4.4). A loop–loop
interaction via the formation of two base pairs triggers the formation of a
duplex of 14 base pairs that links the subunits of the RNA dimer generated
in vitro (Aktar et al. 2013).

Figure 4.4. Dimerization of MPMV gRNA via the Pal SL element. For a
COMMENTARY ON FIGURE 4.4.– The secondary structure of the Pal SL
stem-loop in the gRNA monomer is shown on the left side of the figure. In
blue and black are represented the two Pal SL stem-loops that form a
loop–loop complex, which then leads to the formation of the stable dimer.
4.1.2.2. Dimerization of MMTV gRNA
The gRNA dimer of MMTV, which is present in the virion, has not been
studied by electron microscopy. By analogy with other retroviral species, it
is assumed that the 5' part of the genome is responsible for its dimerization.
In favor of this hypothesis, an in vitro synthesized RNA corresponding to the
5' part of the MMTV genome can form a dimer in vitro (Aktar et al. 2014).
The Pal II element was identified as playing an important role in the
formation of the RNA dimer generated in vitro (see Figure 4.5). RNA
dimerization is decreased if the PBS Pal element is deleted. The action of
this element is rather indirect because it is not a contact point between the
two subunits of the dimer. Although deletion of the Pal II element suppresses
MMTV replication, this has not been shown to be due to impaired gRNA
dimerization (Aktar et al. 2014).
Figure 4.5. Sites involved in the in vitro dimerization of MMTV gRNA. For a

COMMENTARY ON FIGURE 4.5.– The PBS Pal and Pal II elements, which are
upstream of the splice donor site (SD), were identified in vitro using an in
vitro synthesized RNA that corresponds to the 5' end of the MMTV gRNA.
The orange disk represents the 5' cap. Regions and genes are not shown to
scale.
The Pal II element forms a short stem-loop that has a self-complementary
sequence in the apical loop (see Figure 4.6). The apical loop of the stem-loop
is probably directly involved in the intermolecular interaction that links the
subunits of the unstable dimer generated in vitro. This loop–loop interaction
forming a short duplex of six base pairs is thought to initiate the formation of
a long duplex (14 base pairs) linking the subunits of the stable dimer that is
observed in vitro (Aktar et al. 2014).
Figure 4.6. Dimerization of MMTV gRNA via the Pal II element. The secondary
structure of the Pal II stem-loop in the gRNA monomer is shown in the left part of the
figure. Shown in blue and black are the two Pal II stem-loops that link the stable and
unstable dimer subunits via Watson–Crick base pairs. For a color version of this
4.1.3. Structures and mechanisms in deltaretroviruses
Dimerization of deltaretrovirus gRNA has been studied mainly in
HTLV-1. The gRNA dimer of HTLV-1 has not been studied by electron
microscopy. By analogy with other retroviral species, it is assumed that the
5' part of the genome is responsible for its dimerization. In favor of this
hypothesis, an RNA synthesized in vitro and corresponding to the 5' part of
the HTLV-1 genome can form a dimer in vitro. This dimerization is
suppressed by a 32-nucleotide deletion, which is just upstream of the PBS
sequence and contains the sequence called DIS2 (see Figure 4.7) (Greatorex
et al. 1996; Monie et al. 2001). Deletion of this sequence decreases HTLV-1
replication by only 20–25% (Le Blanc et al. 2000). Therefore, the DIS2
sequence identified in vitro may not correspond to the sequence that links

the subunits of the gRNA dimer in the virus particle. In support of this
hypothesis, a recent in vitro study, performed under different experimental
conditions with RNA containing the entire 5'-UTR region and the 5' end of
the gag gene, identified the DIS1 sequence as essential for dimerization (Wu
et al. 2018). In this study, in vitro dimerization was suppressed by mutations
in DIS1 but not by those in DIS2. However, the DIS1 sequence has not been
shown to be required for HTLV-1 replication nor to link subunits of the
RNA dimer generated in vitro or those of the gRNA dimer that is in the
virion.
Figure 4.7. Sites involved in the in vitro dimerization of HTLV-1 gRNA. For a
COMMENTARY ON FIGURE 4.7.– The two DIS elements, which are in the U5
domain downstream of the splice donor site (SD), were identified in vitro
using an in vitro synthesized RNA that corresponds to the 5' end of the
HTLV-1 gRNA. The orange disk represents the 5' cap. Regions and genes
are not shown to scale.
Figure 4.8. Secondary structures of DIS1 and DIS2 sequences in the gRNA
monomer. Shown in blue and black are the two DIS2 sequences that interact
to associate the subunits of the dimer generated in vitro. For a color version of this

The DIS1 and DIS2 sequences each form a short stem-loop (see
Figure 4.8). DIS2 has an apical loop that consists of a single adenine. This is
possible because the CG base pair adjacent to the adenine is unusual. Indeed,
guanine adopts the syn conformation (Monie et al. 2004). The opening of
two DIS2 stem-loops allows the formation of a duplex of 12 base pairs that
associates the subunits of the RNA dimer generated in vitro (Monie et al.
2001).
4.1.4. Structures and mechanisms in gammaretroviruses
Dimerization and encapsidation of gammaretrovirus gRNA have been
studied primarily with MoMuLV. Analysis of gRNA dimers, which were
extracted from immature and mature viral particles, suggests that the
formation of the stable gRNA dimer in the virion depends on the NC that is
released upon cleavage of the Gag precursor by the viral protease (Fu and
Rein 1993). Electron microscopy analysis of the MoMuLV gRNA dimer,
which was extracted from the virion, indicates that the DLS that associates
the two subunits involves less than 50 nucleotides and is located
approximately 466 nucleotides from the 5' end (Murti et al. 1981) (see
Figure 4.9). MoMuLV gRNA dimerization was studied in vitro using in vitro
synthesized RNAs containing the DLS site. In vitro experiments did not
reveal a dimerization site at the DLS (Prats et al. 1990; Tounekti et al. 1992;
Girard et al. 1995). Instead, four contact points (SLA, SLB, SLC and SLD)
associating the two subunits of the RNA dimer generated in vitro were
identified upstream of the DLS (Girard et al. 1995; De Tapia et al. 1998;
Oroudjev et al. 1999; Miyazaki et al. 2010).
Figure 4.9. Sites involved in the dimerization of MoMuLV gRNA. For a
COMMENTARY ON FIGURE 4.9.– The DLS interaction site identified by
electron microscopy is in the encapsidation domain downstream of the splice
donor site (SD). The SLA, SLB, SLC and SLD elements, which are also in the
encapsidation domain and more than 100 nucleotides away from the DLS,

were identified in vitro. The orange disk represents the 5' cap. Regions and
genes are not shown to scale.
Figure 4.10. Secondary structure of a part of the MoMuLV gRNA encapsidation
domain. For a color version of this figure, see www.iste.co.uk/fosse/structures.zip
COMMENTARY ON FIGURE 4.10.– The secondary structure that is shown
corresponds to that present in the gRNA monomer. The self-complementary
sequences are in orange. The UCUG motif in black, which is repeated four
times, is a high-affinity site for the MoMuLV nucleocapsid protein. These
motifs are not accessible to the NC in the gRNA monomer because they are
engaged in base pairs.
The SLA, SLB, SLC and SLD elements in the gRNA monomer
correspond to stem-loop structures (see Figure 4.10) (Tounekti et al. 1992;
Gherghe et al. 2010a). The stem-loops SLA and SLB, which each possess a
self-complementary sequence, play a crucial role in in vitro gRNA
dimerization (Girard et al. 1995; Oroudjev et al. 1999; Ly and Parslow
2002). The gRNA deleted from both stem-loops is unable to dimerize in
vitro and forms a less stable dimer in the virion (Ly and Parslow 2002).
Virus replication is greatly diminished by deletion of the SLA stem-loop but
not the SLB stem-loop (Aagaard et al. 2004; Grohman et al. 2014). Thus,
gRNA dimerization via the SLB stem-loop is not essential for virus viability.
The SLC and SLC stem-loops, whose apical loop has the conserved GACG
sequence, contribute to the stability of the gRNA dimer by being involved in

two loop–loop interactions forming a very short duplex of two base pairs
(Miyazaki et al. 2010).
Figure 4.11. gRNA dimer in the immature virus particle. For a color
corresponds to that of the gRNA in the immature virus particle. The

self-complementary sequences are in orange. The UCUG motif in black that
is repeated four times is a high-affinity site for the MoMuLV NC. The NC
can interact with all four motifs because they are not involved in base pairs.
Figure 4.12. gRNA dimer in the mature virus particle. For a color

corresponds to that of the gRNA in the mature virus particle. The
self-complementary sequences are in orange. The UCUG motif in black that
is repeated four times is a high-affinity site for the MoMuLV NC. The NC
can interact with all four motifs because they are not involved in base pairs.
In vitro studies have probably characterized the sequences that link the
subunits of the gRNA dimer subunits in the virus particle. Indeed, studies
performed using the SHAPE method are in favor of a direct involvement of
the SLA and SLB elements in the association of the gRNA subunits that is
present in both mature and immature virus particles (Gherghe et al. 2010a,
2010b; Grohman et al. 2014). Specifically, the two subunits of the gRNA
dimer, which is present in the immature particle, are primarily associated by
the 10-base pair duplex resulting from the opening and pairing of the two
SLA stem-loops (see Figure 4.11). The dimer is stabilized by two loop–loop
interactions involving the SLC and SLD stem-loops.
The two subunits of the gRNA dimer, which is present in the mature
particle, are associated by the duplex of 10 base pairs SLA–SLA and the
duplex of 16 base pairs resulting from the opening and pairing of the two
SLB stem-loops (see Figure 4.12). NC is likely responsible for the formation
of the SLB–SLB duplex (Girard et al. 1996; Grohman et al. 2014). Both
duplexes confer high stability to the gRNA dimer. In addition, the dimer is
also stabilized by two loop–loop interactions involving the SLC and SLD
stem-loops.
4.1.5. Structures and mechanisms in lentiviruses
4.1.5.1. Dimerization of HIV-2 gRNA
The HIV-2 gRNA dimer has not been studied by electron microscopy. By
analogy with other retroviral species, it is assumed that the 5' part of the
genome is responsible for its dimerization. In favor of this hypothesis, RNAs
synthesized in vitro and containing the 5' part of the HIV-2 genome can form
dimers in vitro. The 5' end of the PBS sequence and the TAR3, PAL and
SL1 elements (see Figure 4.13) have been identified as playing a role in in
vitro dimerization (Dirac et al. 2001; Jossinet et al. 2001; Lanchy et al. 2003;

Purzycka et al. 2011). It has not been determined whether TAR3 and the 5'
end of PBS associate the subunits of the gRNA dimer that is in the mature
and immature viral particles. Mutations designed to prevent dimerization via
the SL1 element have no effect on HIV-2 replication or on the formation of
the gRNA dimer that is in the virion (L’Hernault et al. 2007). However,
analysis of RNA from heterozygous viral particles suggests that the
formation of a six-base pair duplex via two SL1 elements (described below)
is involved in the selective encapsidation of dimeric gRNA (Ni et al. 2011).
Mutations designed to prevent dimerization via the PAL element inhibit
virus replication and significantly decrease the amount of dimeric gRNA in
virions (L’Hernault et al. 2007, 2012).
Figure 4.13. Sites involved in HIV-2 gRNA dimerization. The 5' end of the PBS
sequence and the TAR3, PAL and SL1 elements, which are upstream of SD, were
identified in vitro. The orange disk represents the 5' cap. Regions and genes are not
shown to scale. For a color version of this figure, see www.iste.co.uk/fosse/
structures.zip
At physiological temperature and in the absence of the HIV-2
nucleocapsid protein, an RNA containing the entire 5'-UTR region and the
start of the gag gene forms an unstable dimer in vitro via TAR3 and the 5'
end of PBS and SL1 (Lanchy and Lodmell 2002; Purzycka et al. 2011).
These three elements, two of which are stem-loops (see Figure 4.14),
associate the subunits of the unstable dimer by forming short duplexes
consisting of six to eight base pairs. It has not been determined whether
these elements participate in the formation of an unstable gRNA dimer in the
immature virus particle.
In vitro, at physiological temperature, the conversion of the unstable
dimer into a stable dimer requires the presence of the NC, which, in a first
step, destabilizes the intramolecular base pairing interaction constituted by
the C-box and G-box sequences (Lanchy et al. 2003; Purzycka et al. 2011).
In a second step, the NC, by binding to the upper part of the two SL1
stem-loops, destabilizes the lower part of the latter and thus allows the

release of the two PAL sequences that are partially paired (see Figure 4.15).
In a third step, the PAL sequences pair to form a duplex of 10 base pairs that
are the main contact point of the stable dimer generated in vitro.
Figure 4.14. Folding of the 5' end of HIV-2 gRNA. For a color
COMMENTARY ON FIGURE 4.14.– The folding that is represented
corresponds to that adopted by the monomeric RNA synthesized in vitro and
corresponding to the 5' end of the gRNA. The self-complementary sequences
are in orange (10 nucleotides for the PAL sequence, 8 nucleotides for the
apical loop of TAR3 and 6 nucleotides for the 5' end of the PBS sequence
and the apical loop of SL1). The C-box and G-box sequences (black lines)
form an intramolecular base pairing interaction in the unstable dimer. The
G-box sequence contains the initiation codon of the gag gene.

Figure 4.15. Model for the formation of the stable HIV-2 gRNA
dimer via PAL sequence pairing. For a color version of
COMMENTARY ON FIGURE 4.15.– In this model, two SL1 stem-loops interact
via a loop–loop interaction. This interaction forms a short duplex of six base
pairs, which is one of the contact points associating the subunits of the
unstable dimer. Apical loops with a self-complementary sequence are
represented by a black line. The NC (purple ellipse) binds to the upper parts
of the SL1 stem-loops and opens their lower parts. The opening of the SL1
stem-loop leads the PAL self-complementary sequences (orange line) to form
a duplex of 10 base pairs.
4.1.5.2. Dimerization of HIV-1 gRNA
Dimerization of gRNA could be an early event, as the in vitro transcription
of a DNA corresponding to the 5'UTR-gag region of the proviral DNA
produces dimeric RNA (Darlix et al. 1990). Transcription of proviral DNA
produces three types of unspliced RNAs with one, two or three guanines at the
5' end (Masuda et al. 2015). An RNA synthesized in vitro and corresponding
to the first 344 nucleotides of the HIV-1 genome with two or three guanines at
the 5' end exhibits a weak ability to dimerize, while one with one guanine
dimerizes efficiently (Kharytonchyk et al. 2016). Two or three guanines at the
5' end of the viral genome alter its folding. Interestingly, it is the one-guanine
unspliced RNA that is preferentially encapsidated in the virion as a gRNA
NC
5’
5’
5’ 5’
3’ 3’
3’ 3’ 3’
5’
3’
5’
3’
5’
3’
5’
PAL
PAL
SL1
SL1

dimer (Masuda et al. 2015; Kharytonchyk et al. 2016). Although unstable
dimers can be generated in vitro in the absence of protein, it has not been
shown that gRNA does not require protein to form the immature dimer. An
ex vivo study suggests that the Gag precursor facilitates the formation of the
immature dimer (Chen et al. 2016a).
An ex vivo study performed with retroviral vectors strongly suggests that
dimerization of HIV-1 gRNA occurs prior to its encapsidation into the
immature viral particle (Moore et al. 2007). A study based on total internal
reflection fluorescence microscopy (usually abbreviated TIRF) suggests that
dimeric gRNAs exist at the plasma membrane (Chen et al. 2016a). The
presence of gRNA dimers in the cytosol and at the plasma membrane of
living cells was demonstrated by a more precise study using a form of
super-resolution microscopy called 3D-SIM and ccFCS, which is a form
of fluorescence correlation spectroscopy (Ferrer et al. 2016). The latter study
did not reveal gRNA dimers in the nucleus and is therefore in favor of
gRNA dimerization being initiated in the cytosol; this does not exclude the
formation of dimers at the plasma membrane. Analysis of RNA encapsidated
in viral particles produced by fused cells also suggests that gRNA does not
dimerize in the nucleus (Moore et al. 2009). However, the hypothesis of
gRNA dimerization in the nucleus has not been completely abandoned
because current analytical methods do not allow it to be disproved or
confirmed beyond doubt.
Electron microscopy analysis of the gRNA dimer, which is extracted
from the virion, indicates that the DLS is located within the first 400
nucleotides of the 5' end of the genome and consists of two contact points
(Höglund et al. 1997). The same type of DLS has been observed with atomic
force microscopy with an RNA dimer formed in vitro, with each subunit
corresponding to the first 744 nucleotides of the 5' end of the HIV-1 genome
(Andersen et al. 2004; Pallesen 2011). Results from studies focused on
dimerization of in vitro synthesized RNAs suggest that the TAR and SL1
elements are the two contact points of the DLS observed by both microscopy
techniques (see Figure 4.16).
Dimerization of HIV-1 gRNA has been the subject of numerous in vitro
studies using RNAs containing DLS (see references cited in the study by
Dubois et al. (2018)). The RNA, which is synthesized in vitro and
corresponds to the TAR element, dimerizes efficiently when incubated in the
presence of HIV-1 NC (Andersen et al. 2004). The apical portion of the TAR

stem-loop has a self-complementary sequence (see Figure 4.17) that is
responsible for dimerization.
Figure 4.16. Sites involved in HIV-1 gRNA dimerization. The DLS identified by
electron and atomic force microscopy is thought to involve the TAR and SL1
elements that are upstream of the splice donor site (SD). The orange disk represents
the 5' cap. Regions and genes are not shown to scale. For a color version of this
In the context of a long RNA (744 nucleotides), the TAR element has
only a minor role in NCp7-dependent dimerization in vitro (Andersen et al.
2004). Furthermore, studies of mutants suggest that ex vivo gRNA
dimerization is not dependent on the TAR autocomplementary sequence
(Jalalirad et al. 2012). Thus, the TAR–TAR interaction has not been shown
to be one of the contact points that binds the gRNA dimer subunits in the
virion. The primary and perhaps exclusive role of TAR is its ability to
activate transcription of proviral DNA (Das et al. 2007).
Figure 4.17. Secondary structure of the TAR element. The secondary structure that
is shown corresponds to that present in the gRNA monomer. The self-
complementary sequence (10 nucleotides) initiating dimerization via the TAR
element is in orange. For a color version of this figure, see www.iste.co.uk/fosse/
structures.zip

In the absence of the NC, the SL1 element has been identified as
responsible for the dimerization of an in vitro synthesized RNA
corresponding to the first 615 nucleotides of HIV-1 gRNA (Skripkin et al.
1994). The major role of the SL1 element in in vitro dimerization in the
absence of an NC has been confirmed by other studies (Laughrea and Jetté
1994; Muriaux et al. 1995; Clever et al. 1996). The SL1 element forms a
stem-loop structure, with the apical loop having a self-complementary
sequence of six nucleotides (see Figure 4.18). In many HIV-1 isolates, there
is strong selection pressure to maintain the autocomplementary sequences
GUGCAC and GCGCGC in the apical loop (Hussein et al. 2010). Deletion
of the stem-loop SL1 or mutations in its apical loop, which delete the
autocomplementary sequence, decrease HIV-1 infectivity by a factor of 10 to
1,000 (Paillart et al. 1996b; Clever and Parslow 1997; Laughrea et al. 1997).
Dimerization of gRNA in the cytosol and at the plasma membrane is
significantly decreased by the deletion of the SL1 stem-loop or mutations in
its apical loop (Ferrer et al. 2016). However, the gRNA extracted from
virions is in the form of a dimer when the SL1 stem-loop is deleted or when
its apical loop is mutated (Berkhout and van Wamel 1996; Clever and
Parslow 1997). The gRNA can therefore form ex vivo a dimer that does not
depend on the stem-loop SL1, but this dimer does not allow efficient
replication of the virus. It has been proposed that the SL1 element initiates
the formation of a gRNA dimer that has an optimal conformation for HIV-1
replication (Clever and Parslow 1997).
Under physiological temperature and salinity conditions, and in the
absence of the NC, an RNA corresponding to the 5' part of the HIV-1 gRNA
forms an unstable dimer in vitro via the SL1 stem-loop (Laughrea and Jetté
1996). This dimer may correspond in part to the gRNA dimer that is present
in the immature viral particle. The subunits of this dimer dissociate when
analyzed by agarose gel electrophoresis at room temperature in the absence
of magnesium. The apical loop of the SL1 stem-loop is directly involved in
the loop–loop interaction that binds the subunits of the unstable dimer
(Clever et al. 1996; Paillart et al. 1996a). This interaction forms a short
duplex of six base pairs (see Figure 4.18). Studies by NMR and X-ray
crystallography of a short RNA (23–24 nucleotides) representing the upper
part of the SL1 stem-loop have confirmed the loop–loop interaction (Ennifar
et al. 2001; Kieken et al. 2006).
RNAs synthesized in vitro and representing the 5' end of the HIV-1
genome form stable dimers after incubation at elevated temperatures

(55–65°C) (Laughrea and Jetté 1996; Muriaux et al. 1996a). These in vitro
studies strongly suggest that the subunits of the stable dimers are linked by a
long duplex (28 base pairs) resulting from the opening and pairing of two
SL1 stem-loops (see Figure 4.18). It was shown by NMR that a short RNA
(35 nucleotides) corresponding to the entire SL1 sequence can form the long
duplex (Ulyanov et al. 2006). A subsequent NMR study showed that an
RNA synthesized in vitro and corresponding to the first 344 nucleotides of
the 5' end of the gRNA can form a stable dimer at 37°C whose subunits are
linked by the long duplex (Keane et al. 2016).
Figure 4.18. Dimerization of HIV-1 gRNA via the SL1 element. The secondary
structure of the SL1 stem-loop in the gRNA monomer is shown in the upper part of
the figure. Shown in blue and black are the two SL1 stem-loops that link the stable
and unstable dimer subunits via Watson–Crick base pairs. The internal loops on
which the NC binds are in purple. For a color version of this figure, see
Under physiological conditions, in vitro formation of stable dimers is
strongly stimulated by HIV-1 NC when RNAs contain the SL1 stem-loop
(Muriaux et al. 1996b; Andersen et al. 2004). It is likely that the subunits of
these dimers are associated by the long duplex (see Figure 4.18), but this has
not been demonstrated. The conversion of the loop–loop complex to the long
duplex has been studied in vitro with short RNAs (23 and 35 nucleotides)
and different analytical techniques (gel electrophoresis, fluorescence, mass

spectrometry and NMR) (Rist and Marino 2002; Hagan and Fabris 2007;
Mujeeb et al. 2007). The results of these studies strongly suggest that the
binding of NC on the inner loop destabilizes the apical stem of SL1 without
dissociating the loop–loop complex, thus leading to the formation of the long
duplex. To date, it has not been shown that the long duplex generated in
vitro is the or one of the contact points that associates the subunits of the
gRNA dimer in the mature virus particle.
Figure 4.19. First regulation model of HIV-1 gRNA dimerization by
alternative folding of the 5'-UTR region. For a color version of
COMMENTARY ON FIGURE 4.19.– On the left is the BMH conformation,
which can dimerize because the SL1 element forms a stem-loop. The self-
complementary sequence (orange) is responsible for the loop–loop
interaction in the unstable dimer. Eleven nucleotides of the sequence U5
3’
5’
TAR
.
.
.
A
SL3
PBS
G U
SD
SL2
SL1
3’
5’
TAR
poly(A)
.
.
.
GUA
U5
PBS
SL3
SD

(in red) are paired with the sequence (in green) that contains the initiation
codon (AUG) of the gag gene. These two paired sequences constitute the
beginning and the end of the encapsidation signal Ψ. On the right is shown
the LDI conformation, which cannot dimerize because the SL1 element is
paired in part with the poly(A) and U5 domains.
RNAs corresponding to the 5'-UTR region and the beginning of the gag
gene can adopt two alternative conformations in vitro (Abbink and Berkhout
2003). In the one called BMH, the SL1 element forms the stem-loop
involved in dimerization, while in the other, called LDI, it interacts with the
polyA and U5 elements and thus does not form the stem-loop (see
Figure 4.19). It has been proposed that ex vivo the BMH conformation leads
to gRNA dimerization and inhibits gag gene translation because the
initiation codon of the gag gene is involved in a stable base pairing
interaction that is conserved in different lentiviruses (Abbink and Berkhout
2003; Tran et al. 2015). Mutations that promote BMH folding, however, do
not reduce translation ex vivo (Abbink et al. 2005). The secondary structure
of the gRNA in the virion, which was determined using the SHAPE method
(Watts et al. 2009), is consistent with the BMH conformation. It has been
proposed that ex vivo the LDI conformation is competent for translation of
the gag gene but not for gRNA dimerization. However, LDI folding has not
been characterized ex vivo, and mutations that prevent its formation do not
decrease translation ex vivo (Abbink et al. 2005).
A second alternative folding model of the 5'-UTR region and the
beginning of the gag gene has been proposed from agarose gel
electrophoresis and NMR analyses of an RNA corresponding to the 5' end of
the HIV-1 genome (Lu et al. 2011). In this model, the BMH conformation,
which is slightly different from the first model, is in equilibrium with the
U5-SL1 conformation (see Figure 4.20). This BMH conformation is also
consistent with the secondary structure of the gRNA in the virion, which was
determined using the SHAPE method (Watts et al. 2009). It has been
proposed that gRNA dimerization requires the BMH conformation, while
translation of the gag gene is facilitated by the other conformation. It has not
been shown ex vivo that U5-SL1 folding exists and promotes gag gene
translation.

5’
TAR
poly(A)
PBS
SL3
3’
.
.
.
U5
SL1
SL2
SL4
G
U
A
5’
TAR
poly(A)
A
PBS
SL3
3’
SL1
SL2
.
.
.
U
G
U5
SD
SD
Figure
4.20.
Second
regulation
model
of
HIV-1
gRNA
dimerization
by
alternative
folding
of
the
5'UTR.
For
a
color
version
of
this
figure,
see

COMMENTARY ON FIGURE 4.20.– On the left is shown the BMH
conformation, which can dimerize because the SL1 element forms a
stem-loop. The self-complementary sequence (orange) is responsible for the
loop–loop interaction in the unstable dimer. Eleven nucleotides of the U5
sequence (in red) are paired with the sequence (in green) that contains the
initiation codon (AUG) of the gag gene. These two paired sequences
constitute the beginning and the end of the encapsidation signal Ψ. On the
right is represented the U5-SL1 conformation, which cannot dimerize
because five nucleotides of the autocomplementary sequence (six
nucleotides) are paired with nucleotides of the U5 sequence. The initiation
codon of the gag gene is in the SL4 stem-loop.
4.2. RNA structures and mechanisms regulating gRNA
encapsidation
In different retroviral species, the gRNA is recognized by the Gag
precursor, usually through a specific interaction between the NC domain of
the polypeptide and a structured RNA region that is called the encapsidation
signal or domain. Less than 1% of Gag polypeptides (i.e. about a dozen
molecules) are thought to be involved in gRNA recognition (Jouvenet et al.
2009; Miyazaki et al. 2010). The first studies focused on the identification of
encapsidation signals of different retroviruses were performed with whole
genomes (see references cited in the studies by Jouvenet et al. (2011) and
Maldonado and Parent (2016)). They showed that the 5'-UTR region of
retroviral gRNAs contains a major encapsidation signal. Following these
initial studies, studies concerning the encapsidation of retroviral genomes
were mainly performed with retroviral vectors for a simpler analysis of the
results and a finer characterization of the RNA sequences and structures
involved in gRNA encapsidation.
4.2.1. Structures and mechanisms in alpharetroviruses
Alpharetrovirus gRNA encapsidation has primarily been studied in RSV
(Maldonado and Parent 2016). Although identical, RSV unspliced RNAs are
thought to split into two populations in the nucleus cell (Maldonado and
Parent 2016). The unspliced RNAs in one population serve as genomic

RNAs and interact with one or more Gag polypeptides to pass into the
cytoplasm and then are directed to the plasma membrane where they are
encapsidated into viral particles. It is not known whether it is a monomeric
gRNA or an unstable gRNA dimer that interacts with one or more Gag
precursors. The NC domain of the Gag polypeptide, mainly via its two zinc
fingers, is responsible for the specific encapsidation of RSV gRNA (Méric
and Spahr 1986; Méric et al. 1988). In addition, the formation of the gRNA
dimer in the virion depends on the NC, which is released during the cleavage
of the Gag precursor by the viral protease (Oertle and Spahr 1990; Stewart et
al. 1990).
Figure 4.21. RSV gRNA encapsidation signal. The MΨ encapsidation signal is
upstream of SD and the gag gene. The orange disk represents the cap in 5'. Regions
and genes are not shown to scale. For a color version of this figure, see
The MΨ sequence (160 nucleotides) is the main encapsidation signal of
RSV because it contains much of the information necessary for gRNA
encapsidation and leads to the encapsidation of non-viral RNA when fused
to it. Thus, MΨ creates a 100-fold increase in non-viral RNA encapsidation
(Aronoff et al. 1993; Banks et al. 1999). However, the encapsidation of a
non-viral RNA containing MΨ is decreased by a factor of 3 compared to the
whole gRNA (Banks et al. 1999). There may be a sequence or sequences
downstream of MΨ that serve as secondary encapsidation signals, or that
increase encapsidation by inducing a conformation of the gRNA that is
optimal for the MΨ-Gag precursor interaction. These sequences present in
the gRNA, downstream of the 250th nucleotide of the gag gene, have not
been characterized. As the main RSV encapsidation signal is upstream of
the SD (see Figure 4.21), it should also allow for selective and efficient
encapsidation of viral spliced RNAs. However, analysis of RNAs
encapsidated in virions shows that gRNA encapsidation is 15 times greater
than that of env mRNA (Banks et al. 1999). One hypothesis is incorrect
folding of the MΨ encapsidation signal into the env mRNA due to long-range
interactions between sequences upstream of the SD and those present in
the env gene.

Figure 4.22. Secondary structure of the MΨ encapsidation domain. For a
COMMENTARY ON FIGURE 4.22.– The secondary structure of the MΨ
domain (the dashed boxed portion) corresponds to that present in the RSV
(Pr-C strain) gRNA monomer. Guanine 218 in orange is an important
contact point in the Gag-MΨ interaction.
The secondary structure of the gRNA in the virion and in the cell has not
been determined. However, a recent study determined the secondary
structure of an RNA synthesized in vitro corresponding to the first 636
nucleotides of the RSV genome (Liu et al. 2020). This in vitro study showed
that the MΨ region is highly structured (see Figure 4.22) and that guanine
218 present in the apical loop of the SLC stem plays an important role in
binding the Gag polypeptide to the MΨ domain. The results of this study
combined with those of an NMR study (Zhou et al. 2007) strongly suggest
that guanine 218 interacts with the N-terminal zinc finger of the NC domain
of the Gag precursor. However, the NC domain of the Gag polypeptide has
other contacts with MΨ to package the gRNA. Indeed, mutations in the

apical loop of SLC only moderately decrease viral replication and the
encapsidation of non-viral RNA containing MΨ (Banks and Linial 2000;
Zhou et al. 2007). The MΨ region including the O3 stem and the SLA, SLB
and SLC stem-loops plays a crucial role in encapsidation. Indeed, the
deletion of the L3 stem-loop, which plays an essential role in RNA
dimerization in vitro (see section 4.1.1), did not decrease the encapsidation
of a non-viral RNA containing the other MΨ elements (Banks and Linial
2000).
4.2.2. Structures and mechanisms in betaretroviruses
Betaretroviruses, such as MPMV and MMTV, differ from other retroviral
species in the assembly process of the immature viral particle, which takes
place in the cytoplasm, not at the plasma membrane of the infected cell. It is
during this process that the gRNA is encapsidated into the immature viral
particle through its interaction with several Gag polypeptides. It is not
known whether it is a monomeric gRNA or an unstable gRNA dimer that
interacts with the polypeptides.
4.2.2.1. Encapsidation of MPMV gRNA
The specific gRNA encapsidation of MPMV depends on the C-terminus
of the CA domain (RKK basic motif) and the NC domain of the Gag
polypeptide (Füzik et al. 2016; Dostálková et al. 2018). Although not
demonstrated for MPMV, it is likely that the activity of the NC domain
results in part from its two zinc fingers. However, it has been shown that the
KNKEK basic motif, which is just upstream of the N-terminal zinc finger
(René et al. 2018), plays an important role in encapsidation (Dostálková
et al. 2018).
Figure 4.23. MPMV gRNA encapsidation signal. The encapsidation signal is on
either side of the splice donor site (SD). The two domains that are essential for the
activity of Ψ are in red. The orange disk represents the 5' cap. Regions and genes
are not shown to scale. For a color version of this figure, see www.iste.co.uk/
fosse/structures.zip

The use of retroviral vectors made it possible to identify the region
named Ψ (389 nucleotides), which is the primary signal for MPMV gRNA
encapsidation (Schmidt et al. 2003; Jaballah et al. 2010; Kalloush et al.
2016). The activity of Ψ is primarily due to two domains (see Figure 4.23).
The first domain is just downstream of the PBS sequence and consists of 50
nucleotides. Its 3' end contains the Pal SL sequence that is involved in in
vitro RNA dimerization (see section 4.2.1.1). This sequence is not sufficient
for encapsidation, and it has not been shown to be required for ex vivo
dimerization of gRNA. The second domain is composed of the last 23
nucleotides of the 5'-UTR region and the first 120 nucleotides of the gag
gene. Encapsidation is reduced by 33 times when the first domain and the
first 23 nucleotides of the second domain are deleted (Jaballah et al. 2010).
Encapsidation is reduced threefold when the last 90 nucleotides of the
second domain are deleted (Schmidt et al. 2003). The encapsidation of
spliced RNAs is much less efficient than that of gRNA because they contain
only the first four nucleotides of the second domain of Ψ.
been characterized. The secondary structure of an RNA synthesized in vitro
corresponding to the MPMV encapsidation signal has been determined
(Aktar et al. 2013). This in vitro study showed that the region is highly
structured (see Figure 4.24). The SL3, Gag SL1 and Gag SL2 stem-loops
have minor roles in encapsidation (Kalloush et al. 2019). In contrast, the
encapsidation of an RNA containing Ψ was greatly diminished when the
two long-range interactions (LR-I and LR-II) between the U5 region and
the gag gene were deleted by site-directed mutagenesis (Kalloush et al.
2016, 2019). LR-I and LR-II allow Ψ to adopt a conformation that is
optimal for exposing the Gag polypeptide-interacting sites. In this
conformation, two internal loops, which are accessible and located on either
side of the Pal SL and SL3 stem-loops, constitute two high-affinity sites for
the Gag precursor (Pitchai et al. 2021). These loops are rich in purines and
GU motifs (sequence UUAAAAGUGAAAGUAA for one and sequence
AAGUGU for the other). Since in all retroviruses except HTLV-1, NC binds
preferentially to unpaired guanines and often to the GU motif (see references
in René et al. (2018)), it is likely that on each loop a Gag precursor molecule
binds via its NC domain. Interestingly, the env mRNA, which is only very
weakly encapsidated, does not contain the Gag polypeptide binding site that
is just downstream of the SD. It does have the site upstream of the SD, but

this is partially paired and thus less accessible to the polypeptide (Pitchai
et al. 2021).
Figure 4.24. Folding of the MPMV Ψ encapsidation signal. For a color
COMMENTARY ON FIGURE 4.24.– The folding of Ψ that is shown
corresponds to that present in the MPMV gRNA monomer. The two domains
that are primarily responsible for the activity of Ψ are in green. The two
high-affinity sites for the Gag precursor are dashed. The splice donor site
(SD) is at the 3' end of the SL3 stem-loop. AUG is the initiation codon of the
gag gene.
SL3
SL1
5’
LR-I
3’
LR-II
PBS
G
U
A
Pal SL
SL2
SD

4.2.2.2. Encapsidation of MMTV gRNA
Although not directly demonstrated by mutations targeting the Gag
precursor, it is likely that MMTV gRNA encapsidation is dependent on this
polypeptide. In favor of this hypothesis, mutations in the encapsidation
signal strongly decrease encapsidation and Gag-binding in vitro
(Chameettachal et al. 2021).
The use of retroviral vectors made it possible to identify the region
named Ψ (432 nucleotides), which is the primary signal for MMTV gRNA
encapsidation (Mustafa et al. 2012). This region contains the entire 5'-UTR
domain and at least the first 120 nucleotides of the gag gene (see Figure
4.25).
Figure 4.25. MMTV gRNA encapsidation signal. The encapsidation signal is on
either side of the splice donor site (SD). The orange disk represents the 5' cap.
Regions and genes are not shown to scale. For a color version of this figure, see
The secondary structure of the gRNA in the virion or in the cell has not
been characterized. The secondary structure of an in vitro synthesized RNA
corresponding to the MMTV encapsidation signal has been determined
(Aktar et al. 2014). This in vitro study showed that the region is highly
structured (see Figure 4.26). The long-range LRI interaction between the U5
region and the gag gene likely allows the Ψ to adopt a conformation that is
optimal for exposing the Gag polypeptide-interacting sites. This hypothesis
has not been supported by published results. Strong sites for Gag have not
been identified in vitro in the 3' portion of the Ψ that corresponds to the first
120 nucleotides of the gag gene (Chameettachal et al. 2021). The structure
of the SL4 element, consisting of a stem and two stem-loops, plays a key
role in encapsidation. Indeed, encapsidation is decreased by a factor of 30 to
50 by mutations that suppress this structure (Mustafa et al. 2018). The apical
loop of one of the two stem-loops is composed only of purines
(GGAGAAGAG) and constitutes a strong binding site for the Gag precursor
in vitro. The replacement of these purines by pyrimidines is deleterious to
the encapsidation and binding of Gag to Ψ (Chameettachal et al. 2021). The

sequence consisting of purines is also present in spliced viral RNAs, but
being paired, it is not a high-affinity site for Gag.
The PBS element, which consists of a short stem-loop and an inner loop,
is a second strong binding site for the Gag polypeptide. Mutations in the
PBS element reduce encapsidation by a factor of 20 and significantly
decrease the affinity of Gag for Ψ (Chameettachal et al. 2021). MMTV
spliced RNAs contain the PBS element, which is no more paired than in
gRNA and thus should possess a high-affinity site for Gag. However, it is
not selectively encapsidated in the immature virus particle and weakly binds
the Gag polypeptide (Chameettachal et al. 2021). This could be due to only
one strong site being accessible to Gag (see above) and possibly also due to
the low accessibility of the PBS element in spliced RNAs.
Figure 4.26. Folding of the MMTV Ψ encapsidation signal
SL1
PBS
SL2
SD
5’
3’
SL6
SL5
G
U
A
LRI

COMMENTARY ON FIGURE 4.26.– The folding of Ψ that is shown
corresponds to that present in the MMTV gRNA monomer. The two
high-affinity sites for the Gag precursor are dashed. The splice donor site
(SD) is in the apical loop that binds Gag. Pal II is the primary site of RNA
dimerization in vitro. AUG is the initiation codon of the gag gene.
4.2.3. Structures and mechanisms in deltaretroviruses
The gRNA encapsidation of deltaretroviruses has been studied mainly in
BLV and, as in all retroviral species, it requires the Gag polypeptide.
However, it is interesting to note that in the case of BLV, encapsidation
depends on the NC domain of Gag as well as on its MA domain. The two
zinc fingers of the NC domain, as well as some basic amino acids of the MA
domain, are probably involved in the Gag-signal encapsidation interaction
(Wang et al. 2003). It is not known whether it is a monomeric gRNA or an
unstable gRNA dimer that interacts with the Gag precursor.
Figure 4.27. BLV gRNA encapsidation domains. For a color version
COMMENTARY ON FIGURE 4.27.– The encapsidation signal consists of two
domains (D1 and D2). D1 (147 nucleotides) is located in the 5'-UTR region
downstream of the splice donor site (SD) and extends to the beginning of the
gag gene. D2 (132 nucleotides) is in the central part of the gag gene. The
orange disk represents the 5' cap. Regions and genes are not shown to scale.
The use of retroviral vectors has shown that the encapsidation signal is
discontinuous and composed of two domains that are downstream of the SD
(Mansky et al. 1995). The first domain (D1), which plays the most important
role in encapsidation, starts downstream of the PBS sequence and ends 70
nucleotides after the gag gene initiation codon (see Figure 4.27). The second
domain (D2) is in the part of the gag gene that encodes the CA domain of
the Gag polypeptide. The spliced viral RNAs are barely packaged because
they do not contain D1 and D2. The secondary structure of the 5' end of the
BLV gRNA has not been determined. Use of the mfold software showed

(Mansky and Wisniewski 1998) that both domains have the potential to form
stem-loops (see Figure 4.28). Encapsidation is decreased fourfold by
mutations that do not allow for the formation of a putative stem-loop (SL3)
in D2 (Mansky and Wisniewski 1998). A portion of D1 encompassing the
beginning of the gag gene has the potential to form two stem-loops (SL1 and
SL2) that are largely responsible for encapsidation activity. Indeed,
mutations that prevent the formation of these two stem-loops reduce
encapsidation 40-fold (Mansky and Wisniewski 1998). Encapsidation is
dependent on the sequence that is exposed in each apical loop (Mansky and
Gajary 2002). Encapsidation of HTLV-1 gRNA, which is a deltaretrovirus,
likely involves two stem-loops similar to SL1 and SL2. Indeed, a
BLV-derived RNA can be encapsidated when both the SL1 and SL2
stem-loops of BLV are replaced by those of HTLV-1 (Mansky and Gajary
2002).
Figure 4.28. Stem-loops involved in BLV gRNA encapsidation. Part of the D1
domain can form the SL1 and SL2 stem-loops, while part of the D2 domain can form
the SL3 stem-loop. The initiation codon of the gag gene is in black. For a color
The interaction between the BLV Gag polypeptide and its gRNA has not
been studied. The stem-loop SL3 could interact with the NC domain of Gag
because it contains three unpaired guanines. Indeed, the BLV NC, like the
NCs of other retroviral species, binds preferentially to sequences containing
unpaired guanines. In contrast, HTLV-1 NC does not have this specificity
(Morcock et al. 2002). SL1 and SL2 are probably not high-affinity sites for
the BLV Gag NC domain because they have only one unpaired guanine. The

interaction of Gag with SL1 and SL2 depends primarily on its MA domain.
In favor of this hypothesis, two isolated HTLV-1 stem-loops similar to SL1
and SL2 are two strong sites for HTLV-1 MA protein, while they do not
bind HTLV-1 NC (Wu et al. 2018).
4.2.4. Structures and mechanisms in gammaretroviruses
The encapsidation of gammaretrovirus gRNA has been studied mainly in
MoMuLV and, as in all retroviral species, it requires the Gag polypeptide.
The zinc finger of the NC domain plays a crucial role in the Gag-signal
encapsidation interaction (Gorelick et al. 1988; Berkowitz et al. 1995; Zhang
and Barklis 1995).
Figure 4.29. MoMuLV gRNA encapsidation domain. The encapsidation signal is
located in the 5'-UTR region. It starts at the splice donor site (SD) and ends about 50
nucleotides from the gag gene initiation codon. The orange disk represents the 5'
cap. Regions and genes are not shown to scale. For a color version of this figure,
Several studies strongly suggest that the gRNA that is recognized by Gag
is dimeric (Levin et al. 1974; Méric and Goff 1989; Hibbert et al. 2004). In
vitro results suggest that some of the unspliced RNA molecules may
dimerize during transcription of proviral DNA (Flynn and Telesnitsky 2006;
Maurel et al. 2007). In addition, two ex vivo studies support the formation of
unspliced RNA dimers in the nucleus (Flynn and Telesnitsky 2006; Maurel
and Mougel 2010). The various microscopy techniques and strategies used to
date have not been able to confirm or refute the presence of unspliced RNA
dimers in the nucleus. The Gag polypeptide could be detected in the nucleus
of MoMuLV-infected cells (Nash et al. 1993).
The Ψ region (350 nucleotides), which is located just downstream of the
SD and upstream of the gag gene (see Figure 4.29), is the primary signal for
MoMuLV gRNA encapsidation because its deletion suppresses this process
(Mann et al. 1983). Although the region is sufficient for efficient
encapsidation, the gRNA contains upstream and downstream sequences that
promote optimal encapsidation (see references in the study by D’Souza and
Summers (2005)).

A study conducted with chemical probes and RNA synthesized in vitro
showed that the Ψ region forms an independent domain because its
secondary structure is the same when isolated or present in the RNA
corresponding to the first 725 nucleotides of the MoMuLV genome
(Tounekti et al. 1992). This study suggests that Ψ forms 10 stem-loops in the
gRNA monomer. Interestingly, these include the SLA, SLB, SLC and SLD
stem-loops (see Figure 4.10), which are involved in gRNA dimerization (see
section 4.1.4). Two ex vivo studies performed with retroviral vectors have
shown that the SLC and SLD stem-loops are required for the encapsidation
activity of Ψ (Mougel et al. 1996; Mougel and Barklis 1997).
The UCUG motif, which is present several times in the encapsidation
signal, was identified in vitro as a site with high affinity for the NC
(D’Souza and Summers 2004). Analysis by Weeks’ team of the gRNA
dimer, which is present in the virion, identified two high-affinity regions for
NC in the encapsidation signal (Gherghe et al. 2010b). Each region is
composed of two UCUG motifs (UCUG-UR-UCUG) and is unpaired in the
gRNA dimers (see Figures 4.11 and 4.12). Interestingly, the UCUG-UR-
UCUG sequence is present only in the gRNA encapsidation signal. Like NC,
the Gag precursor binds preferentially to the unpaired UCUG motif
(Gherghe et al. 2010b). The first U and G are the most important in the
UCUG-Gag/NC interaction (Gherghe et al. 2010b). When the guanines of
the four UCUG motifs in the encapsidation signal are replaced by adenines,
the encapsidation of an RNA containing Ψ is decreased by a factor of 100
(Gherghe et al. 2010b). Studies by Weeks’ team converge to propose a
model in which each of the eight UCUG motifs in the immature gRNA
dimer interacts with the NC domain of a Gag polypeptide (Grohman et al.
2014). The conformation adopted by Ψ in the dimeric gRNA is likely the
product of an evolutionary process leading to optimal interactions between
unpaired UCUG motifs and Gag polypeptides.
In vitro studies (Tounekti et al. 1992; D’Souza and Summers 2004;
Gherghe et al. 2010a) suggest that the four UCUG motifs are not accessible
in the gRNA monomer because they are at least partially paired (see
Figure 4.10). Thus, dimerization of the gRNA is likely required for it to
interact with multiple molecules of the Gag precursor. An appealing, but yet
to be proven, hypothesis is that the Gag polypeptide recognizes in the
nucleus the fraction of unspliced dimeric RNAs that thus serve as gRNAs,
while the other fraction of unspliced monomeric RNAs are not recognized

by the polypeptide but serve as mRNAs for the synthesis of Gag and
Gag–Pol precursors.
4.2.5. Structures and mechanisms in lentiviruses
In lentiviruses, such as HIV-1 and HIV-2, the gRNA in the form of a
dimer is encapsidated in the immature virus particle through its interaction
with several Gag polypeptides (Moore et al. 2007; Ni et al. 2011; Chen et al.
2016a; Ferrer et al. 2016).
4.2.5.1. Encapsidation of HIV-2 gRNA
Although this has not been directly demonstrated by mutations targeting
the Gag precursor, it is likely that HIV-2 gRNA encapsidation of HIV-2
gRNA is dependent on this polypeptide. In support of this hypothesis, the
encapsidation of an RNA that corresponds in part to HIV-2 gRNA was
significantly decreased when unpaired guanines, putatively interacting with
the Gag precursor, were replaced by other nucleotides (Umunnakwe et al.
2021).
Three teams have studied HIV-2 gRNA encapsidation by analyzing the
effect of deletions on both sides of the SD. These studies did not lead to a
consensus on the location of the encapsidation signal. Indeed, it has been
localized to the SD either upstream (McCann and Lever 1997; Griffin et al.
2001), downstream (Poeschla et al. 1998) or on both sides (Arya et al. 1998).
been characterized. The secondary structure of an RNA synthesized in vitro
corresponding to the first 560 nucleotides of HIV-2 has been determined
(Purzycka et al. 2011). This in vitro study showed that the 5'-UTR region
and the beginning of the gag gene are highly structured (see Figure 4.30).
Based on the assumption that in the overwhelming majority of retroviral
species, the standard binding site for a Gag molecule is unpaired and
contains at least one guanine, nine putative sites were tested by site-directed
mutagenesis (Umunnakwe et al. 2021). These sites, corresponding to loops,
contain 18 unpaired guanines (see Figure 4.30). Encapsidation is decreased
by a factor of six when all of these guanines are replaced by other
nucleotides. Sites six and nine apparently have no role in encapsidation.

Figure
4.30.
Folding
of
the
5'
end
of
the
unstable
HIV-2
gRNA
dimer.
For
a
color
version
of
this
figure,
see

COMMENTARY ON FIGURE 4.30.– The folding that is shown corresponds to
that of a subunit of the unstable dimer formed by the RNA synthesized in
vitro and corresponding to the 5' end of the gRNA. The two apical loops
linking the subunits of the unstable dimer are in orange. Loop sequences are
indicated where they have been mutated to determine their role in
encapsidation. Guanines that have been replaced by other nucleotides are in
red. With the exception of site #7, which consists of two internal loops, one
site corresponds to a single loop. The C-box and G-box sequences (black
lines) form an intramolecular base pairing interaction in the unstable dimer.
The G-box sequence contains the initiation codon of the gag gene.
In contrast, the other sites are involved in encapsidation and probably
allow the binding of multiple Gag to the immature dimeric gRNA. The
binding of multiple polypeptides to dimeric gRNA produces a synergistic
effect that allows efficient encapsidation. Of the sites that can bind the Gag
polypeptide, site 3 is the most important because replacement of its three
guanines with other nucleotides decreases encapsidation by a factor of about
two (L’Hernault et al. 2007; Baig et al. 2009; Umunnakwe et al. 2021).
4.2.5.2. Encapsidation of HIV-1 gRNA
The specific interaction of the Gag precursor via its two zinc fingers with
the gRNA is necessary for the encapsidation of the latter into the virion
(Aldovini and Young 1990; Gorelick et al. 1990). However, the two zinc
fingers are not equivalent, with the N-terminal playing the most important
role in encapsidation (Gorelick et al. 1993). The immature dimer gRNA that
is present in the cytosol of HIV-1-infected cells (Ferrer et al. 2016) interacts
with a limited number of Gag polypeptide molecules to be transported to the
plasma membrane, where it is incorporated into the assembling virus particle
(Kutluay and Bieniasz 2010; Hendrix et al. 2015). Association of the Gag
polypeptide with unspliced viral RNA could be detected in the nucleus
of HIV-1-infected cells (Ukah et al. 2018; Tuffy et al. 2020), but this
association has not been shown to be the first step in the process of gRNA
encapsidation (Grewe et al. 2012; Tuffy et al. 2020).
Deletions in the region between the SD and the initiation codon of the
gag gene decrease gRNA encapsidation in the virion by at least a factor of
100 (Lever et al. 1989; Aldovini and Young 1990; Clavel and Orenstein
1990). Two studies (Heng et al. 2012; Kharytonchyk et al. 2018) suggest that
the encapsidation signal is the region that lies between the R sequence and

the first 15 nucleotides of the gag gene (see Figure 4.31). In these studies,
the Ψ sequence (246 nucleotides) is sufficient to encapsidate a non-viral
RNA when fused to it, which contains a signal that allows it to be
transported from the nucleus to the cytoplasm. However, another study (Liu
et al. 2017), performed with non-retroviral RNAs also containing a nuclear
export signal, suggests that the domain is not sufficient to allow efficient
encapsidation. This study also suggests that a significant portion (1,061
nucleotides) of the HIV-1 genome consisting of the entire 5'-UTR region and
approximately the 5' half of the gag gene is required to allow optimal
encapsidation of a non-retroviral RNA. Since the identification of the
encapsidation signal was based on the analysis of deleted RNAs, it is likely
that the above studies reached different conclusions because they did not use
the same deletions or the same retroviral vectors. In the study by Hu’s team
(Liu et al. 2017), deletions upstream and downstream of Ψ could change its
folding and thus indirectly inhibit its activity (Das et al. 2012; Kharytonchyk
et al. 2018).
Figure 4.31. HIV-1 gRNA encapsidation domain. The encapsidation signal is located
almost exclusively in the 5'-UTR region. It begins downstream of the R sequence and
ends 12 nucleotides after the gag gene initiation codon. The orange disk represents
the 5' cap. Regions and genes are not shown to scale. For a color version of this
The secondary structure of the gRNA in the virion, which was
determined by the SHAPE method (Watts et al. 2009), is consistent with the
two BMH conformations that have been proposed for the 5'-UTR region (see
Figures 4.19 and 4.20). The Ψ region folds into the form of the U5-AUG
stem and the PBS, SL1, SL2 and SL3 stem-loops. The U5-AUG, SL1 and
SL3 structural elements play an important role in the activity of Ψ, whereas
the PBS and SL2 stem-loops are not directly involved (McBride and
Panganiban 1997; Houzet et al. 2007; Heng et al. 2012).
Using the SHAPE method and an inhibitor that inhibits the binding of NC
to RNA, it was possible to identify seven putative NC sites in the Ψ region
of the dimeric gRNA that is in the virion (Wilkinson et al. 2008). These sites
likely correspond to sites recognized by Gag via its NC domain. The sites

all have one to three unpaired guanines and are downstream of the PBS
stem-loop (see Figure 4.32). The unpaired guanines in the putative Gag sites
were replaced with other nucleotides to study the effect of these substitutions
on encapsidation (Nikolaitchik et al. 2020). The guanines in the apical loop
of SL2 were not mutated because they overlap with the SD. The unpaired
guanines act synergistically in encapsidation. Indeed, encapsidation is
minimally decreased by substitutions located in two putative Gag sites,
whereas it is reduced by a factor of 7 to 13 when four putative Gag sites are
mutated, and by a factor of 34 when all putative Gag sites are mutated except
the one containing the SD. Of the six sites studied, the two that are located
in the lower part of the SL1 stem-loop are the most important for
encapsidation. Interestingly, these two sites interact strongly with the Gag
precursor in vitro (Abd El-Wahab et al. 2014). Taken together, these results
suggest that binding a Gag polypeptide to each of the seven sites produces a
synergistic effect that allows for optimal encapsidation of immature dimeric
gRNA.
Figure 4.32. Secondary structure of a Ψ portion of HIV-1. For a color

corresponds to a portion of Ψ in one subunit of the stable dimer that is
present in the virion. The seven putative Gag sites are in black, and they are
annotated I, II and III according to their strength (strong, medium and
weak). The PBS stem-loop upstream of annotated site II is not shown, nor is
the SL1–SL1 interaction linking the two dimer subunits. Only the lower part
of the SL1 stem-loop, containing two putative strong binding sites for Gag, is
visible in this secondary structure model. The initiation codon of the gag
gene is boxed.

Conclusion
As shown in this book through its four chapters and the literature cited,
the study of structure–function relationships of retroviral RNAs has been
energized over the last three decades by research on HIV-1, which is the
main causative agent of the AIDS pandemic. During this period, new
molecular biology technologies based on retroviral vectors, imaging and the
SHAPE method, among others, have made it possible to determine ex vivo
and in vitro the secondary structures of RNAs associated with specific
functions. These structures are distributed in the coding and non-coding
regions of the retroviral genome. The 5'-UTR non-coding regions of
retroviruses contain the most secondary structures with functional roles.
The retroviral 5'-UTR is highly structured, and some of its structural
elements perform different functions at different stages of the retroviral
cycle. The Psi domain of MoMuLV, which is in the 5'-UTR and consists of
four stem-loops, is a prominent example. Indeed, this domain has three
functions: nuclear export of unspliced RNA, dimerization and encapsidation
of gRNA. The secondary structures of the entire 5'-UTR regions of several
retroviruses have been well-characterized, but the three-dimensional
structures have not. Structural studies based on improvements of existing
tools (NMR and cryo-electron microscopy) and technological innovations
are needed to determine the tertiary structure of a 5'-UTR region at good
resolution. To elucidate the molecular basis of a function associated with a
5'-UTR region during a specific step of the retroviral cycle, structural
analysis of the 5'-UTR region will have to be performed in the presence of
the viral/cellular proteins required for that step. In addition, the folding
dynamics of RNA regions and their potential alternative folds must be taken
into account to accurately determine the structure–function relationships.

Structure-dependent functions are based on RNA–RNA (tRNA–PBS,
gRNA dimerization), DNA–RNA (first strand transfer and strand transfers
responsible for recombination events) and RNA–protein (TAR–Tat,
RRE–Rev and Psi–Gag in the case of HIV-1) interactions. The secondary
structures responsible for these functions are generally involved in
intermolecular interactions via unpaired and accessible nucleotides, such as
the loop–loop interaction in gRNA dimerization, the TAR–Tat interaction in
HIV-1 proviral DNA transcription and the Psi–Gag interaction in gRNA
encapsidation.
Although a wealth of knowledge about the structure–function
relationships of retroviral RNAs has been accumulated over the past three
decades, much remains to be discovered about the RNA-dependent
molecular mechanisms that regulate several steps of the retroviral cycle. To
address the emergence of antiretroviral resistance, it is important to have
several families of compounds that inhibit HIV-1 replication. To date, there
is no clinically used antiretroviral that inhibits HIV-1 replication by
specifically interacting with an RNA structure with a function. Further
research and knowledge are therefore required to design such an
antiretroviral.

Glossary
Alternative splicing: the splicing that produces different mRNAs from the
same pre-messenger RNA.
Amino acid: an organic compound possessing a carboxyl group (–COOH)
and a primary (–NH2) or secondary (–NH–) amine group. Amino acids are
the basic constituents of proteins.
Apical loop: a loop at the end of a stem-loop.
Branch point: a conserved sequence in the intron that contains an adenosine
and plays an essential role in splicing.
Bulge: a loop on only one of the two strands of a stem.
Cap: methylated guanosine in position N7 that is linked to the first
nucleotide of the mRNA by a 5'–5' triphosphate linkage. This is necessary
for the protection of mRNA and its translation by the ribosome.
Chemical probe: this is a chemical agent that enables the identification of
paired and unpaired nucleotides in the nucleic acid studied. The
identification of the nucleotides modified by the chemical agent is performed
by primer extension by means of a reverse transcriptase (indirect method) or
by the use of RNA marked at one end (direct method).
Cis-regulatory element: a stimulatory or inhibitory sequence that acts on
the DNA or RNA molecule that contains it.

Clinical isolate: an isolate from a patient.
Coaxial stacking: a process in which two RNA helices form a contiguous
helix that is stabilized by the stacking of bases at the interface of the two
helices.
Complex retroviruses: retroviruses whose genome also codes for accessory
and regulatory proteins.
Crm1 pathway: the nuclear export pathway requiring the Crm1 protein, also
called exportin 1 (XPO1). The activity of Crm1 depends on its association
with Ran-GTP, which is a small GTP-bound G protein. Crm1–RanGTP
transports a very large number of proteins from the nucleus to the cytoplasm
by recognizing the NES they possess.
Encapsidation: the process by which the retrovirus genomic RNA is
incorporated into the virus particle.
Enzymatic probe: an enzyme (nuclease) that allows us to identify the paired
and unpaired nucleotides in the accessible regions of a nucleic acid. The
identification of the enzyme cleavage sites is done by primer extension by
means of a reverse transcriptase (indirect method) or through the use of
RNA marked at one end (direct method).
Ex vivo: experiments performed on cells in culture.
Exon: the part of the gene that is retained after splicing. It is usually coding.
hnRNP proteins: proteins classified into different groups (A, H, etc.) that
are present in heterogeneous ribonucleoprotein particles consisting of
proteins and pre-messenger RNA. They are involved in alternative splicing.
In vitro: experiments performed in test tubes to reproduce biological
processes.
In vivo: experiments performed on a living organism.
Internal loop: a loop that connects two stems.
Intron: the part of the gene that is not retained after splicing. It is generally
non-coding.

Glossary 119
Isolate: a virus isolated from a host and well-characterized genetically. It
reproduces in cell culture.
Kilobase: the unit of measurement corresponding to 1,000 nucleotide bases.
Loop: a sequence of unpaired nucleotides in an RNA secondary structure.
Multiple loop: a loop that connects three or more stems.
NXF1/NXT1 pathway: the main pathway of nuclear export of cellular
mRNAs involving the heterodimer formed by the proteins NXF1 (also called
Tap) and NXT1 (also called p15).
Oncogene: a gene whose expression promotes carcinogenesis.
Primer: a short strand of DNA or RNA complementary to a part of a
template that a DNA polymerase uses to start DNA synthesis by copying this
template.
Secondary structure: the secondary structure of a single-stranded
DNA/RNA molecule describes base pairings (Watson–Crick and G-U type)
and unpaired bases. A secondary structure consists of stems, stem-loops,
bulges, apical loops, inner loops and multiple loops. Numerous secondary
structures of various RNAs have been constructed from the experimental
data obtained with the chemical and enzymatic probes.
SHAPE: a method that makes it possible to determine the secondary
structure of RNA. It is based on a chemical agent that acylates the 2'-OH
group of a nucleotide when the latter is flexible; in general, this corresponds
to an unpaired nucleotide. The identification of nucleotides modified by the
acylating agent is performed by primer extension using a reverse
transcriptase.
Simple retroviruses: retroviruses whose genome codes only for structural
proteins and the three viral enzymes (IN, PR and RT).
Spliceosome: a large complex of proteins and small nuclear RNAs that is
responsible for splicing pre-messenger RNAs.

Splicing: the maturation process of the pre-messenger RNA that eliminates
introns and joins exons. It is based on the reactions involving cutting and
ligation of the pre-messenger RNA strand.
SR proteins: proteins rich in arginines and lysines that bind to
pre-messenger RNAs and are required for various steps of the spliceosome
assembly.
Stem: a paired region that forms a helix.
Stem-loop: a hairpin structure consisting of a stem with one end closed by a
loop called an apical loop.
Template: a strand of nucleic acid that is copied by a polymerase.
Tertiary interaction: a long-distance interaction that participates in the
formation of the three-dimensional structure of RNA.
UBA: a protein domain consisting of about 40 amino acids that binds
ubiquitin. This domain has been proposed to limit ubiquitin chain elongation
and to direct ubiquitinated proteins to degradation via the 26S proteasome.
Virion: a mature and infectious viral particle.

List of Acronyms
3D-SIM three-dimensional super-resolution structured illumination
microscopy
AIDS acquired immunodeficiency syndrome
ALV avian leukosis virus
BIV bovine immunodeficiency virus
CA capsid protein
CAE cytoplasmic accumulation element
ccFCS cross-correlation fluctuation correlation spectroscopy
cDNA complementary DNA
Crm1 chromosome region maintenance 1
CRS cis-active repressive sequences
CTE constitutive transport element
DIS dimer initiation site
DLS dimer linkage structure
DNA deoxyribonucleic acid

DR direct repeat
EIAV equine infectious anemia virus
eIF4E eukaryotic translation initiation factor 4e
F-MLV Friend murine leukemia virus
gRNA genomic RNA (of the retrovirus)
HIV-1 human immunodeficiency virus type 1
HIV-2 human immunodeficiency virus type 2
hnRNP A1 heterogeneous nuclear ribonucleoprotein A1
hRIP human Rev-interacting protein
HTLV-1 human T-cell leukemia virus type 1
HTLV-2 human T-cell leukemia virus type 2
Hu Hu antigen R
IN integrase
IRES internal ribosome entry site
ITAF IRES trans-acting factors
JDV Jembrana disease virus
JSRV Jaagsiekte sheep retrovirus
kb kilobase
L leader (the sequence between the PBS and the gag gene)
LRR leucine-rich repeat
LTR long terminal repeat

List of Acronyms 123
MA matrix protein
MLV murine leukemia virus
MMTV mouse mammary tumor virus
MoMuLV Moloney murine leukemia virus
MPMV Mason–Pfizer monkey virus
mRNA messenger RNA
NC nucleocapsid protein
NES nuclear export signal
NLS nuclear localization signal
NMR nuclear magnetic resonance
NPC nuclear pore complex
NRS negative regulator of splicing
NTF2 nuclear transport factor 2
NTF2L NTF2-like domain
NXF1 nuclear RNA export factor 1
NXT1 NTF2-related export protein 1
PBS primer binding site
PPT polypurine tract
PR viral protease
Psi packaging signal
PTB polypyrimidine tract-binding protein

PTE post-transcriptional element
R repeat (the repeated sequence in the 5' and 3' of the retroviral
genome)
Rev regulator of expression of virion proteins
REV-A reticuloendotheliosis virus type A
RmRE RNA element required for Rem responsiveness
RNA ribonucleic acid
RRE Rev responsive element
RRM RNA recognition motif
RSL R region stem-loop
RSV Rous sarcoma virus
RT reverse transcriptase
RxRE Rex responsive element
SAXS small-angle X-ray scattering
SHAPE selective 2'-hydroxylacylation analyzed by primer
extension
SIV simian immunodeficiency virus
SL stem-loop
SRV-1 simian retrovirus type 1
ssDNA strong-stop DNA
SU surface protein
Tap Tip-associated protein

List of Acronyms 125
TAR transactivator response element
Tat transactivator of transcription
TIM-TAM Tat IRES modulator of Tat mRNA
TM transmembrane protein
TRBP TAR RNA binding protein
TREX1 transcription–export complex 1
tRNA transfer RNA
U2AF U2 auxiliary factor
U3 unique 3' sequence of the retroviral genome
U5 unique 5' sequence of the retroviral genome
UBA ubiquitin-associated domain
UPF1 up-frameshift 1
UTR untranslated region

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Index
A, B, C
acquired immune deficiency
syndrome (AIDS), 1, 115
bulge, 33, 34, 37, 56
chaperone, 11, 29, 76
chemical probes, 40, 107
D, E
decapsidation, 7
dimer, 12, 57, 60, 73, 74, 76–79, 81,
85, 86, 89, 91, 97, 99, 104, 108,
110
duplex, 16, 17, 19, 21, 22, 25, 76, 77,
79, 81, 83, 85–88, 91, 92
encapsidation, 10, 50, 73, 74, 81, 82,
86, 89, 94, 96–108, 110–112, 115,
116
export, 8, 9, 44–46, 48–51, 53, 54,
56, 57, 111, 115
G, H
Gag precursor, 10, 17, 73, 74, 81, 89,
96–98, 100, 108, 110
genomic RNA (gRNA), 2, 9, 12, 16,
22, 26, 39, 50, 72–74, 77–79, 81,
85, 86, 88, 89, 91, 96, 97, 99, 100,
102–104, 106, 108, 111
host cell, 5, 7
I, L
initiation, 13, 14, 17, 19, 25, 34, 36,
60–63, 65–69, 71, 72, 87, 94, 96,
101, 104–106, 110, 111, 113
intermolecular, 16, 116
intramolecular, 16, 42, 66
loop
apical, 28, 33, 35, 75, 82, 91, 99,
102, 105, 112
inner, 46, 48, 93, 103
N, O
nuclear magnetic resonance (NMR),
16, 22, 91, 93, 98
nucleocapsid (NC) protein, 4–6, 11,
12, 16–19, 21–26, 28–30, 73, 74,
76, 81, 82, 84–86, 88, 89, 91–93,
96–100, 104–107, 111
oncogene, 39

P, R
primer, 14, 19
binding site (PBS), 2, 13, 15–19,
21, 68, 71, 78, 79, 85–87, 100,
103, 104, 111–113, 116
proviral DNA, 9, 33, 34, 88
retroviral vectors, 50, 51, 89, 100,
102, 104, 107, 111
reverse transcriptase (RT), 1, 4, 5, 11,
13, 14, 18, 19, 21, 26, 28–30, 39
ribosome, 2, 60, 71, 72
S, T
secondary structure, 11, 17, 19, 22,
42, 57, 94, 98, 100, 102, 108
SHAPE, 17, 22, 59, 85, 94, 111, 115
splicing, 9, 39, 41, 42, 71
stem-loop, 22, 25, 28, 33–35, 46, 54,
68, 69, 71, 75, 81, 82, 86, 91, 94,
99, 102, 105, 107, 112
strand transfer, 21, 26, 28, 29
template, 28, 29
transcription, 3, 7–9, 11–14, 19, 21,
26, 28, 31, 33–38, 54, 56, 68, 73,
88, 90, 106, 116
reverse, 3, 7, 11, 12, 73
translation, 51, 60–62, 65–69, 94
tRNA, 5, 14, 16
U, V, X
unspliced mRNA, 4, 39, 50, 63, 64,
67, 69
virion, 5–10, 14, 17, 73, 74, 77, 78,
80–82, 86, 88–90, 94, 97, 98, 100,
102, 107, 108, 110, 111, 113
X-ray crystallography, 47, 59, 91

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