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Scherer et al., Sci. Adv. 8, eabl4895 (2022) 7 January 2022
SCIENCE ADVANCES | RESEARCH ARTICLE
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CORONAVIRUS
SARS-CoV-2 nucleocapsid protein adheres to replication
organelles before viral assembly at the Golgi/ERGIC
and lysosome-mediated egress
Katharina M. Scherer1†, Luca Mascheroni1†, George W. Carnell2, Lucia C. S. Wunderlich1,
Stanislaw Makarchuk3, Marius Brockhoff1, Ioanna Mela1, Ana Fernandez-Villegas1,
Max Barysevich1, Hazel Stewart4, Maria Suau Sans2‡, Charlotte L. George2, Jacob R. Lamb1,
Gabriele S. Kaminski-Schierle1, Jonathan L. Heeney2, Clemens F. Kaminski1*
Despite being the target of extensive research efforts due to the COVID-19 (coronavirus disease 2019) pandemic,
relatively little is known about the dynamics of severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2)
replication within cells. We investigate and characterize the tightly orchestrated virus assembly by visualizing
the spatiotemporal dynamics of the four structural SARS-CoV-2 proteins at high resolution. The nucleoprotein
is expressed first and accumulates around folded endoplasmic reticulum (ER) membranes in convoluted layers
that contain viral RNA replication foci. We find that, of the three transmembrane proteins, the membrane pro-
tein appears at the Golgi apparatus/ER-to-Golgi intermediate compartment before the spike and envelope
proteins. Relocation of a lysosome marker toward the assembly compartment and its detection in transport
vesicles of viral proteins confirm an important role of lysosomes in SARS-CoV-2 egress. These data provide
insights into the spatiotemporal regulation of SARS-CoV-2 assembly and refine the current understanding of
SARS-CoV-2 replication.
INTRODUCTION
Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) is
an RNA virus and the causative agent of coronavirus disease 2019
(COVID-19) (1). To date, more than 263 million cases of this dis-
ease have been diagnosed, resulting in more than 5.4 million deaths
(2). Great efforts have been made in the development of measures
for containing the spread of SARS-CoV-2, including the repurposing
of previously produced drugs (3), therapies (4), and the development
of vaccines (5).
While new diagnosis, prevention, and treatment options for
COVID-19 continue to emerge at a rapid pace, the understanding of
the biology of SARS-CoV-2 advances more slowly. Unraveling the
mechanisms of transmission and replication of this virus is crucial
for the development of rationally designed drugs and vaccines, and
to understand the long-term effects of the disease, allowing re-
searchers to develop countermeasures against evolving SARS-CoV-2
variants of concern.
SARS-CoV-2 spreads among humans primarily via respiratory
droplets when two individuals are in close proximity (6). It is an
enveloped virus that enters the cells of the respiratory tract through
the interaction of the receptor binding domain on the spike (S) pro-
tein and the angiotensin-converting enzyme 2 receptor on the cell
surface (7). The positive sense, single-stranded RNA genome of
SARS-CoV-2 is then released into the host cell cytosol and is directly
translated. Two large open reading frames (ORF1a and ORF1ab) are
translated into large polyprotein complexes (pp1a and pp1ab), which
are cotranslationally and posttranslationally cleaved to generate
16 nonstructural proteins, for which characterization is ongoing (8).
The remaining ORFs encode the four structural proteins of SARS-
CoV-2 (9). In coronaviruses in general, the nucleocapsid (N) pro-
tein encapsulates the viral RNA (9,10), the S protein mediates cell
entry (7), the membrane (M) protein is embedded in the envelope
and thought to provide a scaffold for viral assembly (11), and the
envelope (E) protein forms ion-conductive channels in the lipid viral
envelope (12). Upon infection by SARS-CoV-2, the virus initiates
the biogenesis of replication organelles (ROs) containing inter-
connected perinuclear double-membrane structures such as double-
membrane vesicles (DMVs), which are derived from, and tethered
to, the endoplasmic reticulum (ER) (13). It is assumed that these
structures protect the viral RNA from degradation by cellular ribo-
nucleases and detection by host cellular immune sensors during ge-
nome replication (14). This hypothesis has been corroborated by the
recent finding of Klein etal., who have proven the presence of viral
RNA inside DMVs (15,16). The DMVs have a pore in their double-
membrane lining, by which the RNA is thought to access the cytosol
(17). The assembly of mature SARS-CoV-2 virions occurs within the
ER-to-Golgi intermediate compartment (ERGIC) (8,13). The egress
of coronaviruses is assumed to occur via exocytosis (15). Recent evi-
dence suggests that newly formed SARS-CoV-2 virions reach the cell
periphery using lysosome trafficking (18).
The interactions of each SARS-CoV-2 protein with a series of
host cell proteins have been partially studied by combining light
microscopy and proteomics (19,20). Gordon etal. (19) used confo-
cal microscopy to study the distribution of two of the structural pro-
teins of SARS-CoV-2in infected Caco-2 cells at one time point after
infection. The imaging revealed a cytosolic signal for the N protein
and strong colocalization of the M protein with the Golgi apparatus.
However, the location of the viral proteins during infection is very
1Department of Chemical Engineering and Biotechnology, University of Cambridge,
Cambridge, UK. 2Department of Veterinary Medicine, University of Cambridge,
Cambridge, UK. 3UK Dementia Research Institute, Cambridge, UK. 4Division of
Virology, Department of Pathology, University of Cambridge, Cambridge, UK.
*Corresponding author. Email: cfk23@cam.ac.uk
†These authors contributed equally to this work.
‡Present address: London School of Hygiene and Tropical Medicine, Keppel St,
London WC1E 7HT, UK.
Copyright © 2022
The Authors, some
rights reserved;
exclusive licensee
American Association
for the Advancement
of Science. No claim to
original U.S. Government
Works. Distributed
under a Creative
Commons Attribution
License 4.0 (CC BY).
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dynamic and complex because it is highly dependent on their inter-
action with each other and the host cell, conditions that cannot be
simulated by transfection. In the current work, we add to the under-
standing of SARS-CoV-2 replication by studying the timing and
location of the interplay between all four structural proteins of
SARS-CoV-2 and the host cell over the time course of the infec-
tion cycle.
The replication of SARS-CoV-2 is known to extensively change
the localization and reshape the morphology of cell organelles and
the cytoskeleton within the host cell. Such morphological alterations
have recently been studied in Calu-3 cells using both optical and
electron microscopy (13). The study by Cortese etal. analyzed
infected cells at a series of time points after infection to detail the
progression of the viral cycle, focusing on the host cell structures in
detail. They demonstrated the progressive fragmentation of the Golgi
apparatus, the recruitment of peroxisomes to the sites of viral repli-
cation, and the reshaping of the vimentin network to accommodate
the DMVs. While electron microscopy highlighted cellular structures
with high definition, the viral proteins were visualized with lower
resolution using confocal microscopy. The power of super-resolution
optical microscopy has been demonstrated by the application of
three-dimensional (3D) STED (stimulated depleted emission micros-
copy) to reveal the formation of a vimentin cage around ROs. How-
ever, the latter technique cannot be performed in high-throughput
fashion and is not easily adapted for multiplexed imaging of several
proteins simultaneously.
Here, we use a range of light microscopy techniques, particularly
a combination of wide-field, confocal, light sheet, and expansion
microscopy, to overcome some of these limitations. To obtain high-
quality imaging data, Vero cells were used for infection because their
morphology is well suited for fluorescence imaging and single cell
analysis. In addition, numerous SARS-CoV-2 studies based on Vero
cells exist, allowing us to put our results into context. For this study,
a fixation protocol that permits transport of infected Vero cells from
class 3 containment laboratories to high-resolution imaging facilities
was developed. Establishing immunostaining protocols for the im-
aging of multiple SARS-CoV-2 and host cell proteins simultaneously
provided well- defined and controlled snapshots in up two four colors
at subwavelength resolution at different infection stages. We present
a detailed investigation of the spatiotemporal organization of the four
structural proteins of SARS-CoV-2 within the host cell during an in-
fection cycle. Specifically, we focus on expression kinetics, the dy-
namic location at different host cell compartments during assembly
and egress as well as organelle and cytoskeleton rearrangement that
is associated with these processes. The four structural proteins are
expressed differentially. On the basis of the expression patterns, clas-
sification criteria are defined and three distinct infection stages are
identified. Sorting of single cells in these distinct infection stages is
used to assess the dynamics of host cell remodeling. We find that
reshaping of microtubules, relocation of lysosomes, and fragmentation
of the Golgi apparatus largely correlate with the local accumulation
of the three viral transmembrane proteins, S, E, and M proteins. By
combining expansion microscopy and light sheet microscopy, we
have produced volumetric maps of protein distributions in whole in-
fected cells. In particular, we see that the N protein associates with
convoluted and fused membrane compartments. The N protein
accumulates in the outer layers of those compartments that fold
around ER membranes and contain at least one double-stranded
RNA (dsRNA) focus, suggesting that they are viral ROs (vROs).
RESULTS
The cellular distribution of SARS-CoV-2 structural proteins is
tightly regulated in space and in time
We first optimized cellular fixation and staining protocols, using
transfection to express the four structural proteins of SARS-CoV-2
individually in Vero cells. We also optimized fixation (formaldehyde
and glyoxal) and permeabilization (Triton X-100 and saponin) re-
agents. Each cellular structure has its own ideal immunostaining
conditions (fig. S1); the ER was best fixed in a glyoxal buffer, pre-
serving the fine structure of the tubular regions. In contrast, the Golgi
apparatus was only stained when fixed with formaldehyde inde-
pendently of the detergent. As a final example, lysosomal staining
was only achieved after permeabilization with saponin. The optimal
staining conditions for the cellular structures being investigated de-
termined the choice of experimental conditions for each sample. A
summary of the optimized fixation and permeabilization conditions for
each of the structures investigated in this work is presented in table S1.
The immunostaining of the transfected cells with the selected an-
tibodies was successful in all fixation and permeabilization condi-
tions tested (fig. S2). The pattern of the S protein staining was
different in the two fixation conditions (formaldehyde and glyoxal).
However, we did not note any differences when fixing and staining
infected cells in these two conditions. It has been previously observed
that the intracellular localization of viral proteins can differ con-
siderably when comparing an individually expressed viral protein
and the same protein within an infected cell (19). These observa-
tions confirmed that investigations should be carried out in virus-
infected cells.
In this study, we fixed infected cells at multiple time points after
infection (5, 7.5, 10, 12, and 24 hours). The spatial distribution of
the viral proteins changed over time and varied between individual
cells at the same time after infection, particularly at later time points.
A high degree of cell-to-cell variability in infection is expected and
has been observed for a range of mammalian viruses. Known sources
for this variability can be the difference in the number of virions
that cells encounter, genetic variation of virus particles, and the state
of the host cell during infection. However, we identified similarities
in the expression and distribution of the viral proteins in individual
cells within the heterogeneous population and across time points.
These patterns correspond to distinct events in the replication cycle.
We used these patterns to classify the cells into different categories.
This analysis on single cells rather than the population average proves
useful as it allows us to gain a clearer picture of how the virus cycle
is staged in time and to connect certain steps in the viral replication
cycle with morphological changes in the host cell.
We found that three different categories or stages were sufficient
to classify the status of any cell in the population (Fig.1A). At an
early infection stage [5 hours post-infection (hpi)], cells were seen
to express the N protein only. At this stage (referred to as stage 1),
the N protein is not homogeneously distributed inside the host
cell cytosol but forms small puncta. From 7.5 hpi onward, the other
structural proteins of SARS-CoV-2—S, M, and E proteins—are ex-
pressed and localized in a compact juxtanuclear membrane com-
partment (stage 2). Cells that were characterized by fragmentation
and spreading of the compartments containing the S, M, and E pro-
teins were classified as stage 3; in these cells, the N protein is highly
dispersed in the cytoplasm. At stage 3 (abundantly present from 10 hpi
onward), we observed small dots of all four structural viral proteins
at the plasma membrane, indicating the formation and trafficking
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Fig. 1. The cellular distribution of structural proteins of SARS-CoV-2 is tightly regulated in space and in time. (A) Three categories describing different cell states
were identified on the basis of the distribution pattern of the N and S proteins during the late phase of the SARS-CoV-2 replication cycle. These categories were termed
stages 1, 2, and 3, respectively. For each stage, a representative confocal microscopy image is shown. Blue: DAPI (4′,6-diamidino-2-phenylindole)–stained nuclei; magenta:
N protein; green: S protein. Scale bar, 10 m. (B) Colocalization between SARS-CoV-2 N, M, and S proteins. Cells were fixed with glyoxal and permeabilized with saponin.
Left: Representative confocal images of infected Vero cells in infection stages 2 and 3. Blue: DAPI-stained nuclei; magenta: N protein; cyan: M protein; green: S protein.
Scale bar, 5 m. Right: Colocalization between viral proteins N, S, and M at different infection stages determined using the Spearman coefficient method (stage 1: n = 20;
stage 2: n = 25; stage 3: n = 20). Manders coefficients are shown in fig. S4A. (C) Left: Representative confocal images of SARS-CoV-2–infected Vero cells in infection stages
2 and 3. Cells were fixed with formaldehyde and permeabilized with Triton X-100. Blue: DAPI-stained nuclei; magenta: N protein; green: S protein; cyan: E protein. Scale
bar, 5 m. Right: Colocalization between viral proteins N, S, and E at different infection stages determined using the Spearman coefficient method (stage 1: n = 16; stage 2:
n = 19; stage 3: n = 18). Manders coefficients are shown in fig. S4B. Significance was tested for all datasets with an unpaired Mann-Whitney test. See figs. S5 and S6 for more
exemplary images.
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of mature SARS-CoV-2 virions. These relative timings within the
replication cycle were not previously known in this detail. The findings
were enabled by the classification strategy described here, which
considers the staging of cells individually rather than population
averages at different times after infection.
The selection of antibodies against the structural proteins allowed
us to image three of the four structural proteins at once. The M and
E protein could not be visualized simultaneously because the re-
spective antibodies belonged to the same host species. Instead, we
immunostained two separate sets of samples, costaining either for
N, M, and S proteins (Fig.1B) or for N, E, and S proteins (Fig.1C).
We expected the M and E proteins to show a similar pattern of
accumulation in the same host cell membrane compartment as the
S protein, as these are all transmembrane proteins. Representative
images and quantitative colocalization analysis confirmed that M
(Fig.1B) and E proteins (Fig.1C) co-occurred foremost with the
S protein and not the N protein. Consequently, M and E pro-
teins follow the same accumulation and fragmentation pattern as
the S protein.
By comparing the colocalization values between N and trans-
membrane proteins (M, S, and E proteins) within the two separate
sample sets for stages 2 and 3, we noticed that, whereas the Spearman
coefficients (see the “Colocalization analysis” section) were close to
zero in the first sample set (Fig.1B), they were increased in the sec-
ond sample set (Fig.1C). This is not due to a different localization
pattern of the proteins in the separate sample sets but due to a dif-
ferent capability to visualize the viral proteins depending on the per-
meabilization reagents used for immunostaining. The use of strong
(Triton X-100) instead of mild (saponin) detergents was required to
visualize the N protein at the juxtanuclear membrane compartment,
in addition to the bright N protein puncta, where the transmembrane
proteins also localize. The stronger detergent also leads to an increased
staining of cytoplasmic, dispersed N protein (fig. S3). The colocal-
ization of N protein with the three transmembrane proteins at this
compartment is in line with the current model for SARS-CoV-2 as-
sembly, where viral nucleocapsids are trafficked to membrane com-
partments enriched with M, S, and E proteins for assembly.
The kinetic profile of SARS-CoV-2 replication
To assess SARS-CoV-2 replication kinetics, we determined the frac-
tion of cells expressing each of the four structural proteins and the
fraction of cells in which dsRNA was present (indicating initiation of
viral RNA transcription) at each time point (Fig.2A). At 5 hpi, 5 to
10% of cells were positively stained for dsRNA and N protein, but
none of the other structural proteins. Consistently, we detected
released viral transcripts in the cell supernatant by quantitative
real-time transcription polymerase chain reaction (RT-qPCR) from
5 hpi onward (Fig.2B). These data confirm the observations of
Cortese et al. in Calu-3 cells, where PCR, an infectivity assay, and
transmission electron microscopy (TEM) were used. In the latter re-
port, the release of viral RNA and infectious virus was observed in
parallel with the appearance of DMVs at 6hpi under similar experi-
mental conditions (13). This suggests that the timing of events
during viral replication is comparable between Calu-3 and Vero cells.
At 7.5 hpi, we observed that the fraction of cells positive for N protein
increased by up to ~15%, with about a third of the cells also express-
ing the other three structural proteins. From 10 hpi on, infected cells
were expressing all four structural proteins at similar levels. This is
again consistent with a significantly increased infectious titer at 10 hpi
Fig. 2. Stepwise expression of the four structural SARS-CoV-2 proteins and
dsRNA correlates with staged release of viral transcripts (5 hpi) and infectious
virus (7.5 to 10 hpi), respectively. (A) The fraction of cells positive for each of the
structural proteins of SARS-CoV-2 or viral dsRNA was determined by immunostaining
of infected Vero cells. For each time point, 30 to 35 wide-field microscope images,
corresponding to 1000 to 1500 cells per sample, were analyzed. For the control
time point of 0 hpi, around 250 cells were analyzed per sample. For the count
of N- and S-positive cells, three samples per time point were analyzed. (B) The
copy number of the viral transcripts in the cell supernatant was measured by
RT-qPCR. Release of viral RNA was observed from 5 hpi onward when first cells
started expressing N protein. (C) The infectious titer of the cell supernatant was
determined by the plaque assay. Newly formed infective SARS-CoV-2 virions were
released from cells from 10 hpi onward. For both assays, two replicates were
carried out. Significance was tested with an unpaired t test. (D) Stage-dependent
replication kinetics. ns, not significant.
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(Fig.2C), confirming the completion of the replication cycle and the
production of new viruses. For most cells, it appeared that M and
E proteins were expressed simultaneously with the S protein. How-
ever, in a few cells, only fluorescence signal from the M, but not the
S, protein was detected (~5% of infected cells; see representative cell
in fig. S7). In contrast, E and S proteins always occurred together. This
indicates that the M protein is expressed before S and E proteins. At
24 hpi, we observed a doubling in the number of infected cells compared
to 12 hpi. We then tracked the average expression level over time by
measuring the average fluorescence intensity per cell (fig. S8). For all
four viral proteins, the trend was similar: The average expression levels
per cell increased until 12 hpi when they saturated. While the average
values at 12 and at 24 hpi are comparable, we note that the distributions
of values are more homogeneous at 24 hpi than they are at 12 hpi.
Last, we classified the cells, according to them being in three dif-
ferent stages, to quantify the kinetic profile of the infection process
(Fig.2D). Between 5 and 10 hpi, we saw a strong shift from stage 1
to stages 2 and 3. At 7.5 hpi, 50% of cells express all four structural
proteins, with an equal number of cells observed in stages 2 and 3
(compact versus fragmented juxtanuclear membrane compartment).
From 10 hpi on, we observe a rapid increase in cells with fragmented
compartments (stage 3) that are dominating the population of in-
fected cells (~75%), whereas the fractions of cells in stages 1 and 2
remain low. This indicates that the transition from stage 2 to stage 3
(compact to fragmented juxtanuclear membrane compartment) mainly
occurs between 7.5 and 10 hpi. This transition also coincides with a
significantly increased production of mature virions at 10 hpi (Fig.2C).
The N protein accumulates around folded ER membranes
in convoluted layers that connect to viral RNA
replication foci
As shown in Fig.1, the intracellular location of the N protein is
distinct from that of the other three structural proteins: Initially, the
N protein accumulates exclusively in small puncta; as the infection
progresses, cytosolic signal gradually increases alongside the puncta
(Fig.3A). In parallel, the number, as well as the size, of N protein
puncta grow significantly (fig. S9). At closer inspection of the larger
puncta in images of infected cells fixed at 10 hpi, the round struc-
tures were found to be shaped like vesicles with an outer layer
containing N protein and a hollow center (Fig.3B, left image). We
propose that these N protein layers are formed at the vROs.
To find support for this hypothesis, we applied expansion
microscopy (21) to investigate these structures in better detail. This
super-resolution technique provides an at least fourfold increase in
resolution via a 64× volumetric expansion of the sample. In these
higher resolved images of the N puncta, we detected that several of
the N compartments consisted of double layers of the protein (Fig.3B,
right image, and movie S1). In addition, we observed that single small
compartments were often fused to larger convoluted 3D structures
(Fig.3C and movies S2 and S3).
The inner N protein compartments measured ~275nm on average
in diameter (Fig.3D). vROs contain single DMVs and DMV packets
(vesicle packets) (15). DMVs formed by SARS-CoV-2 contained in
the ROs are about 300nm in diameter (13,15), which is in agree-
ment with the structures presented. It is accepted that the DMVs
formed by coronaviruses are used by the virus as a protective environ-
ment for replication of its RNA genome (14,15), and the presence of
SARS-CoV-2 RNA in such structures was recently verified by elec-
tron microscopy (15).
It has been established that ROs are derived from ER membranes
and serve as an anchor for the viral replication and transcription
complex (RTC) (22). dsRNA is considered a viral replication inter-
mediate, indicating the proximity of RTCs. By costaining the N protein,
ER, and dsRNA, and then acquiring confocal images of nonexpanded
cells (fig. S10, A and B) as well as expanded cells (Fig.3E), we found
that, at all stages of infection, the N protein–containing compartments
were always associated with the ER. Moreover, the ER membranes
seemed clustered at the spots where those compartments are present.
In the expanded samples, we observed that the N protein formed
layers around the highly convoluted ER membranes (Fig.3F). This
was observed for single small (<1 m), larger fused (>1 m), as well
as double-layered N protein compartments.
Analogously to the N protein–enriched compartments, the dsRNA
foci were also always associated with the ER (Fig.3E). Only some of the
N protein–enriched compartments seemed to colocalize with dsRNA
foci, whereas many replication foci were not co- occurring with the
N protein compartments. A quantitative analysis of confocal images
of nonexpanded cells showed that the fraction of closely associated
compartments and foci decreased for cells in later infection stages
(fig. S11), when the number of dsRNA foci increased (fig. S10B).
However, single-image cell sections might be misleading, as they
omit information on either side of the focal plane. To analyze the
connection between dsRNA foci and N protein–layered compart-
ments, we acquired volume sections of 13 expanded cells using light
sheet microscopy (movie S4). In most samples, most dsRNA foci are
located in a region immediately adjacent to the nucleus. We further
noted that most N protein compartments were connected to at least
one RNA replication focus, which was usually situated in the outer
layer of the compartment (Fig.3G).
The S protein accumulates in Golgi/ERGIC compartments
and transport vesicles containing a lysosome marker
Next, we aimed to determine with which organelles the SARS-CoV-2
transmembrane proteins directly interact during assembly and egress.
Because of the selection and limitation of the used antibodies, we
could only visualize S protein simultaneously with the host cell
structures. However, the three transmembrane proteins S, M, and E
show a high degree of colocalization, making it likely that they
behave in a similar fashion.
The S protein was seen to be at least partially located in the Golgi
apparatus and ERGIC from a co-occurrence with the respective or-
ganelle markers 130 kDa cis-Golgi matrix protein (GM130) and lectin
mannose protein 1 (LMAN-1) during stages 2 and 3 (Fig.4,AandB).
This finding corresponds with observations made previously also for
SARS-CoV-1 (23). We quantified this colocalization by determining
the Spearman coefficients, which exhibited average values of ~0in
control cells and in cells at stage 1 but increased significantly at stages 2
and 3in all cases.
Concurrent to the enrichment of S protein at Golgi and ERGIC
membranes at stage 2, small spots of S protein appeared in the cyto-
plasm (Figs.1A and 4A). This indicates trafficking of the S protein,
and supposedly also M and E proteins, away from the Golgi and
ERGIC membranes. These diffraction-limited spots could either be
transport vesicles containing viral proteins in their lipid membranes
or newly formed virions. It has recently been reported that lyso-
somes are used by the virus to exit the cell and that mature virions
exploit this for transport to the cell surface (18). When we stained
the cells for the lysosome marker lysosome-associated membrane
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Fig. 3. The SARS-CoV-2 N protein is organized in layered structures that are strongly interwoven with the topology of the ER and contain RNA replication foci.
(A) Confocal microscopy cannot resolve the substructure of N protein puncta in the cytosol of infected Vero cells (blue: nuclei; magenta: N protein). Scale bar, 10 m.
(B) The combination with expansion microscopy reveals double layers of N protein within the larger compartments (red arrow). Scale bars, 1 m. Linear expansion factor
of 4.2. (C) Combining expansion with light sheet microscopy reveals the convoluted nature of the N protein structures in 3D. Dotted lines outline the boundaries of each
convoluted compartment in the maximum intensity projection. Scale bar, 2 m. Expansion factor of 4.2. (D) The inner circular layer of the N protein double-layer compart-
ments has an average diameter of 275 nm. Thirty-eight compartments from 10 cells at 12 hpi were analyzed. The micrograph size is 1.25 m per side. (E) Representative
confocal image of an expanded, infected Vero cell stained for the nucleus (blue), ER (calnexin; orange), N protein (magenta), and dsRNA (cyan). Scale bar, 5 m. Expansion
factor of 4.2. (F) The N protein (magenta) forms layers around ER membranes (yellow). Scale bars, 1 m. Expansion factor of 4.2. (G) A combination of expansion and light
sheet microscopy reveals that dsRNA foci (cyan) sit in the layers of the N compartments (magenta). Scale bar, 5 m. Expansion factor of 4.2. The size of the smaller micro-
graphs is 5 m per side.
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Fig. 4. The S protein accumulates at the Golgi apparatus as well as the ERGIC and colocalizes with lysosomes during late infection stages. (A) Representative
confocal images of SARS-CoV-2–infected Vero cells in infection stages 2 and 3 stained for nuclei (blue), N protein (magenta), S protein (green), and Golgi apparatus
(GM130; cyan). Scale bar, 20 m. See fig. S12 for more exemplary images. Colocalization analysis shows partial spatial correlation between S protein and the Golgi appa-
ratus when S protein expression is detected (from stage 2 onward). Control: n = 111; stage 1: n = 20; stage 2: n = 21; stage 3: n = 59. See fig. S13A for Manders coefficients.
(B) Representative confocal images of SARS-CoV-2–infected Vero cells in infection stages 2 and 3 stained for nuclei (blue), N protein (magenta), S protein (green), and
ERGIC (LMAN-1; cyan). Scale bar, 20 m. See fig. S14 for more exemplary images. Partial spatial correlation between S protein and the ERGIC is detected from stage 2 on-
ward. Control: n = 44; stage 1: n = 6; stage 2: n = 41; stage 3: n = 58. See fig. S13B for Manders coefficients. (C) Representative confocal images of SARS-CoV-2–infected Vero
cells stained for S protein (green) and lysosomes (LAMP1; yellow). Scale bar, 5 m. Spot-to-spot distance analysis reveals increased fraction of co-occurring S protein and
lysosome spots at stage 3 compared to stage 2. Significance was tested with an unpaired t test with Welch’s correction for unequal SDs.
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glycoprotein 1 (LAMP1), we found that the spots containing S pro-
tein were often also positive for LAMP1 (Fig.4C). We quantified
co-occurrence of S protein and LAMP1 for stages 2 and 3 using a
spot-to-spot distance analysis. When the centers of spots in both
channels were within a distance of 280 nm, they were considered as
co-occurring. The fraction of co-occurring spots increased from ~20
to ~50% for both S protein and lysosomes at stage 3. These findings
confirm that lysosomes can be used for the shuttling of virions,
further supporting the role of lysosomes in SARS-CoV-2 egress.
The infection alters the morphology and location of host cell
organelles and cytoskeleton
We further analyzed the morphological changes of the host cell
organelles involved in SARS-CoV-2 assembly and egress as well as
the cytoskeleton at different stages of infection. The most notable
morphological change that we noted was a fragmentation of the
Golgi compartment. To quantify this fragmentation, we measured
the angle spanned by the Golgi apparatus around the nucleus, as
depicted in Fig.5A. In cells with fragmented compartments, the an-
gles were typically larger than 180° and often close to 360°. Thus, we
distinguished between cells with a compact (<180°) or fragmented
(>180°) Golgi compartment. The histograms represent the distribu-
tions of angles measured in the cell population at different times after
infection. At 5 hpi, we observed almost no fragmentation. At 12 and
24 hpi, however, the fraction of cells with fragmented Golgi compart-
ments was increased. This corresponds to cells in late infection stages
(stage 3), when new mature viruses were being produced and released.
We also noticed that the lysosomes undergo a spatial redistribu-
tion during SARS-CoV-2 infection (Fig.5B). At stage 1, the lyso-
somes were larger on average than in control cells (control: mean
area=0.94 m2, n=56; stage 1: mean area=1.22 m2, n=21) but
were still homogeneously distributed within the cytoplasm. When
cells started to express S, M, and E proteins (stage 2), the lysosome
marker LAMP1 was recruited to the Golgi/ERGIC compartments
containing the three viral transmembrane proteins, likely through
fusion of the lysosomes with the Golgi and ERGIC membranes. The
fragmentation of the Golgi apparatus (stage 3) corresponded to a
spread of membrane fragments enriched with viral proteins and
LAMP1in the cytoplasm. The correlation of the lysosome signal with
the viral proteins N and S (measured via the Spearman coefficient
after Otsu thresholding) was moderate at stages 2 and 3 (Fig.5B).
There are two distinct sources from which the lysosome signal orig-
inates: small and bright compartments resembling the typical lyso-
some shape (Fig.5C) and LAMP1 accumulated at the Golgi and ERGIC
membranes (Fig.5D), but with markedly lower intensity. We used
manual thresholding to filter out the weaker signal and only investi-
gate the correlation between the small, bright lysosomal compartments
with the viral proteins. We detected no correlation between S protein
and the lysosomes in that case. Furthermore, there was a negative
correlation between N and the lysosomes at stage 3. At this stage, the
N protein was widely distributed in the cytoplasm but excluded from
the location of the lysosomes. This indicates that the correlation
between the viral proteins and LAMP1 only occurs at the compart-
ments after a mixing of membranes.
SARS-CoV-2 infection leads to a remodeling of the microtubule
network. Through a directionality analysis, we found that, from
stage 2 onward, the network loses its orientation (Fig.5E). In non-
infected cells, microtubules spread from the microtubule-organizing
center close to the Golgi apparatus to the extremities of the cell. In
late infection stages, the microtubule filaments were absent from the
juxtanuclear area and they were entangled when compared to con-
trol cells. For cells in stages 2 and 3, we also detected a loss of cell
stiffness, which we measured using atomic force microscopy (AFM;
fig. S17). This could be driven by a remodeling of the actin network,
which is regarded as the overriding, although not the sole, determi-
nant of cell stiffness (24).
DISCUSSION
We applied advanced fluorescence microscopy to investigate the
expression kinetics and spatial arrangement of the four structural
SARS-CoV-2 proteins and studied their colocalization with host cell
compartments in detail. We observed that the expression of the
structural proteins of the virus is tightly staged, with notable differ-
ences between N and the three transmembrane proteins. The N protein
accumulates mainly in small foci that grow in size and number during
the course of infection. Sample expansion in combination with light
sheet microscopy revealed that single N protein compartments com-
prise layered structures of N protein. The compartments resemble
complex and convoluted 3D structures as might result from the fu-
sion and engulfment of smaller vesicular subunits. We believe them
to be part of the ROs formed by SARS-CoV-2. There are several
indicators to support this notion. First, the shapes of the N protein
compartments resemble those of ROs investigated by electron
microscopy, where interconnected DMVs and vesicle packets were
observed (15). Second, the smallest structural units we could identi-
fy within these convoluted structures were vesicles whose average
size was ~275 nm, which matches the size reported for DMVs in
Vero cells (15). Third, it is known that coronaviruses remodel the
host cell ER membranes to integrate the vROs (25,26). We found
that the N protein–containing compartments are tethered to ER
membranes. Last, we reasoned that if the N protein was associated
to the vROs, the N compartments would be associated with the viral
RTCs. We detected the RTCs by staining of dsRNA, an intermediate
of viral RNA replication. It needs to be mentioned that the amount
of genomic viral RNA that is present in infected cells is under-
estimated by using a dsRNA antibody that indicates only replication
factories with high dsRNA content (27). Through volumetric imaging,
we confirmed that at least one dsRNA focus is usually associated with
the outer layer of an N protein compartment, which might consist of
several fused sections. We also noted that, similarly to the N protein
compartments, the dsRNA foci are always connected to the ER network.
For viral genomic RNA compared to dsRNA, a stronger colocal-
ization with the N protein would be expected. Using in situ hybrid-
ization assays for visualizing viral genomic RNA, Stertz etal. (16) for
SARS-CoV and Lee etal. (27) for SARS-CoV-2 could indeed demon-
strate a high co-occurrence of N protein and viral RNA in infected cells.
Because one of the functions of the N protein is the encapsula-
tion of the viral RNA, its presence at/around the DMVs and colocal-
ization with proteins forming the RTCs would not be unexpected in
accordance with previous reports for SARS-CoV (16). However, to
our knowledge, association of N protein to ROs has not been reported
before. But it seems to make sense from the virus perspective since
such an arrangement would spatially connect replication and nucleo-
capsid formation.
If we assume that the N protein–containing structures are indeed
part of the vROs, the question remains where exactly the N protein
is located within those compartments and how this association is
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Fig. 5. SARS-CoV-2 infection alters the morphology and location of organelles and the cytoskeleton in Vero cells. (A) The angular distribution of Golgi compart-
ments around the nucleus was used to distinguish between cells with compact (angle < 180°) and fragmented Golgi apparatus (angle > 180°). The fraction of cells with
fragmented Golgi apparatus increased over the time course of infection. The fragmentation of the Golgi apparatus upon SARS-CoV-2 infection was used to sort cells into
infection stage 2 or 3 (Fig. 2D). (B) Left: Representative confocal images of lysosomes in infected Vero cells at different infection stages. Blue: DAPI-stained nuclei; magenta:
N protein; green: S protein; yellow: lysosomes (LAMP1 staining). Scale bar, 10 m. See fig. S15 for more exemplary images. Right: Colocalization of the lysosome marker
LAMP1 with the N and S proteins. (C) Otsu thresholding allows correlation between all LAMP1 and SARS-CoV-2 protein signal. (D) Manual thresholding, however, filters
out the weaker LAMP1 signal such that only bright, small lysosomal compartments are taken into account (stage 1: n = 25; stage 2: n = 34; stage 3: n = 39). (E) Left: Repre-
sentative confocal images of microtubules in control (mock infected) and infected Vero cells (stages 1, 2, and 3). Scale bar, 10 m. See fig. S16 for more exemplary images.
Right: The directionality coefficient was calculated for subareas of the microtubule network. Each data point corresponds to one subregion inside a cell. Changes in the
network because of SARS-CoV-2 infection lead to significant reduction in directionality at stage 1, which is even more pronounced at stages 2 and 3 when all four structural
proteins are expressed. Significance was tested with a Mann-Whitney test.
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formed. It is possible that the protein accumulates in the inter-
membrane space of the DMV envelope. Another possibility is the
accumulation of the N protein at ER membranes while or after they
are reshaped into ROs. Nonstructural proteins of coronaviruses are
known to reshape host cell membranes to induce the formation of
DMVs (14). Either a specific interaction with one or more SARS-
CoV-2 nonstructural proteins or a curvature-driven binding mech-
anism could drive an accumulation of N protein. In both cases,
accumulation might be affected by a propensity of N protein to
phase-separate with RNA (28–33). It has been proposed that N protein
plays a dual role: The unmodified protein forms a structured oligomer
suitable for nucleocapsid assembly, while the phosphorylated protein
forms a liquid-like compartment for viral genome processing (34).
For both processes, association of N protein to the ROs would thus
be beneficial.
A study based on cryo–electron tomography (ET) showed that
strands of naked viral RNA are located within the DMVs (25), which
are thought to leave the DMVs through a pore in the membrane lining
(17). However, it is currently not known where and when the newly
synthesized viral RNA is encapsidated by the N protein. In the light
of the data presented here, we speculate that association of the viral
RNA and the N protein to form viral ribonucleocapsid protein com-
plexes (vRNPs) occurs at the membrane of the vROs. This process
might occur either before or in concurrence with the release of the
newly synthesized RNA into the cytosol. In this sense, we interpret
the increasing cytosolic signal of the N protein in late infection stages
as an accumulation of vRNPs in the cytosol before and during virus
assembly.
Using multicolor imaging and colocalization analysis, we show
that the SARS-CoV-2 S, M, and E proteins all localize at the Golgi
and ERGIC compartments (identified by protein markers) in agree-
ment with previous reports (8,13). These compartments are also thought
to be places of viral assembly (35). Using in situ cryo-ET, formation
of SARS-CoV-2 virus particles has been observed in regions with a
high vesicle density and close to ER- and Golgi-like membrane
arrangements (15). In addition, vesicles containing assembled virus
particles as well as budding events at and into those vesicles were
furthermore captured by cryo–FIB/SEM (focused ion beam/scanning
electron microscopy) and high-resolution cryo-ET (36). This fits well
with the picture of assembly that results from our light microscopy
data. Accumulation of the N protein at the perinuclear compartment
upon expression of S, M, and E strongly indicates these membranes
as points of assembly.
Our study showed furthermore that M protein is recruited to this
area slightly earlier than S and E proteins, suggesting a predominant
role of M protein for controlling the spatial organization of the
transmembrane proteins and initiating the assembly of SARS-CoV-2.
For other coronaviruses, it has indeed been shown that interactions
between M proteins form a lattice into which the other two trans-
membrane proteins of the virus are incorporated (37–39). More-
over, we detected that the N protein partially accumulates at the
Golgi region, but only after the expression of the other three
structural proteins has taken place. It is known that the assembly of
coronaviruses is dependent on the M and E proteins, and for SARS-
CoV also on the N protein (39). In particular, the carboxyl tail of the
SARS-CoV-1 M protein interacts specifically with the N protein (40).
Our results suggest that, also for SARS-CoV-2, the M protein is
responsible for the recruitment of the N protein to the Golgi/ERGIC
membranes.
At late stages of infection, we detected an enrichment of the lyso-
somal protein LAMP1 at the membrane compartments together
with the structural transmembrane proteins of SARS-CoV-2. This
suggests a mixing of membranes or a shift/modification in the endo-
lysosomal transport pathways. Immediately after, diffraction-limited
spots of S protein can be seen in the cytoplasm. These spots often
co-occur with the lysosome marker. Our results corroborate the re-
cent finding that lysosomes are used by coronaviruses for their cell
egress. Ghosh etal. (18) have observed vesicles containing single
SARS-CoV-2 virions with hallmarks of lysosomes using TEM. It is
not clear whether the fluorescent spots we observe consist only of
vesicles enriched with viral transmembrane proteins or mature virions.
Nevertheless, the co-occurrence of viral proteins and lysosome
markers indicates an immediate onset of the egress pathway after
the expression of the M, S, and E proteins. However, we detected an
increase from ~20 to ~50% co-occurrence at transition from a com-
pact (stage 2) to a fragmented Golgi apparatus (stage 3), indicating a
surge in viral egress. It is not clear what causes the fragmentation of
the Golgi apparatus. It might be caused by an indirect toxic effect
due to the accumulation of viral proteins by merging of lysosomes
with Golgi membranes and/or the manipulation of the microtubule
network, which plays an important role in shaping Golgi structure
and function (41). We found support for the latter by measuring a
remarkable rearrangement of the microtubule network after expres-
sion of the three SARS-CoV-2 transmembrane proteins, but before
Golgi fragmentation occurs. However, further work is needed to
elucidate which factors contribute to the defect in the organization
of the Golgi compartments. We envisage that the methods presented
in this study could furthermore be used for studying the role of the
nonstructural proteins of SARS-CoV-2, the kinetics of the viral
genome replication, as well as the relationship between the viral RNA,
the N protein, and the vROs.
MATERIALS AND METHODS
Biosafety
SARS-CoV-2 infection of cells was conducted at containment level 3.
Inactivation of SARS-CoV-2 through fixation was validated using
previously published protocols (42). The results of this experiment
were reviewed and approved by the biosafety committee of the De-
partment of Chemical Engineering of the University of Cambridge.
Chemicals
Methanol-free formaldehyde was purchased from Thermo Fisher
Scientific; the ampoules were used immediately after opening, and
any leftover formaldehyde was discarded. Glyoxal (40% in water)
was purchased from Sigma-Aldrich; the glyoxal solution was heated
and mixed before use to solubilize precipitated glyoxal. Saponin,
Triton X-100, and ammonium chloride were purchased from
Sigma-Aldrich. All chemicals used for sample expansion (glutaral-
dehyde, 50% in water, sodium acrylate, N,N′-methylenbisacrylamide,
and acrylamide) were purchased from Sigma-Aldrich and used as
received. Lyophilized proteinase K was purchased from Thermo
Fisher Scientific. Atto 590–conjugated phalloidin was purchased
from Sigma-Aldrich and solubilized in methanol according to the
manufacturer’s instructions.
Antibodies
All the antibodies used in this study are reported in Tables1and2.
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Cells and viruses
Vero cells (American Type Culture Collection, CCL-81) were cul-
tured under standard conditions (37°C and 5% CO2) in Dulbecco’s
minimum essential medium (DMEM) (Sigma-Aldrich) supplemented
with 10% heat-inactivated fetal bovine serum (Gibco), antibiotics/
antimycotics [penicillin (100 U/ml), streptomycin (100 g/ml), and
Gibco amphotericin B (0.025 g/ml) Gibco], and 2 mM l-glutamine
(GlutaMAX, Gibco). Cells were cultured in T-75 polystyrene flasks;
splitting took place when cultures reached ~80% cell confluency.
For all experiments, cells below passage number 20 were used.
The BetaCoV/Australia/VIC01/2020 strain of SARS-CoV-2 was
obtained from the Victorian Infectious Diseases Reference Laboratory,
Melbourne (43), through Public Health England. This virus was
passaged once in Vero cells for stocks used in this study. The virus
was titrated in standard six-well plaque format on Vero cells, and one
batch of virus was used for all experiments. Virus sequences were
verified by deep sequencing.
Transfection of Vero cells
The four structural proteins of SARS-CoV-2 were expressed in Vero
cells using a pEVAC vector backbone. The day before transfection,
Vero cells were seeded at 30% confluence in eight-well Ibidi -slides
(catalog no. 80826) in antibiotic-free medium. Cells were transfected
with Lipofectamine 3000 transfection reagent (Thermo Fisher Sci-
entific) using 100ng of plasmid DNA and 0.3l of Lipofectamine
reagent per well. Cells were incubated for 48 hours under standard
conditions before fixation and immunostaining as detailed below.
Infection of Vero cells
The day before infection, Vero cells were seeded at 60% confluence
in 24-well plates equipped with 13-mm round glass coverslips (VWR,
catalog no. 631-0150). Cells were washed once with phosphate-buffered
saline (PBS) before incubation with BetaCoV/Australia/VIC01/2020
diluted in PBS at a multiplicity of infection of 5. Incubation took place
at room temperature (RT) on a rocking plate for 1 hour, whereupon
inocula were removed and cells were washed twice with PBS and
replenished with complete DMEM. Infection was allowed to progress
under standard conditions (37°C, 5% CO2) for 0-, 5-, 7.5-, 10-, 12-,
and 24-hour time periods. Cells were fixed with either formaldehyde
(4% methanol-free formaldehyde in 100 mM cacodylate buffer) or
glyoxal [4% glyoxal and 10% ethanol in acetate buffer (pH 5), as
previously reported (44)] after the removal of spent media. Fixation
was carried out at 37°C for 20min.
Plaque assay
Plaque assays were performed as previously described for SARS-CoV-1,
with minor amendments (45,46). The day before infection, Vero cells
were seeded at 30% confluence in six-well plates. These subconfluent
monolayers were infected with 10-fold serial dilutions of each sample
in duplicate, diluted in serum-free media, for 1 hour at RT on a
rocking plate. After the removal of the inocula and washing with PBS,
3ml of 0.2% agarose in virus growth media was overlaid, and the cells
were incubated at 37°C for 72hours. At this time, the overlay media
were removed, and cells were washed with PBS and fixed overnight
with 10% formalin. Fixed monolayers were stained with toluidine
blue, and the plaques were counted manually.
Polymerase chain reaction
The viral load of the media collected before cell fixation at 0-, 5-,
7.5-, 10-, 12-, and 24-hour time points after infection was measured
and quantified via RT-qPCR. Total RNA extraction of the media
was performed using the QIAamp Viral RNA Mini Kit (QIAGEN)
following the manufacturer’s instructions. Five microliters of the
RNA extraction final elution was reverse-transcribed to cDNA and
amplified according to the manufacturer’s protocol using the TaqMan
Fast Virus 1-Step Master Mix (Thermo Fisher Scientific). The primer
pair was as follows: 5′CAGGTATATGCGCTAGTTATCAGAC-3′
(forward) and 5′-CCAAGTGACATAGTGTAGGCAATG-3′ (re-
verse). The probe used was as follows: 5′-[6FAM]AGACTAAT-
TCTCCTCGGCGGGCACG[TAM]-3′ (Sigma-Aldrich). Analysis
was performed using the Rotor-Gene 6000 Series Software 1.7
(Corbett Life Sciences, QIAGEN).
To generate RNA standards for qRT-PCR, a 97-nucleotide frag-
ment of the spike ORF was cloned into the pJET1.2 vector (Invitrogen).
Following linearization with Hind III, invitro RNA transcripts were
generated using the T7 RiboMAX Express Large Scale RNA Produc-
tion System (Promega). Transcripts were purified (RNA Clean and
Concentrator, Zymo Research) and the integrity was confirmed by
gel electrophoresis.
Immunostaining of fixed cells
Cells were fixed as detailed in the “Infection of Vero cells” section.
The choice of fixative was determined by the structures to be immuno-
stained in each sample, as detailed in table S1. A summary of the
fixations and permeabilization conditions for the micrographs shown
in this paper is reported in table S2. Fixed cells were incubated with
50 mM NH4Cl in PBS for 10min to quench fixation. Cells were
permeabilized with either 0.2% saponin or 0.2% Triton X-100 (see
table S2) in PBS for 15min and then blocked with 10% goat serum
(Abcam) in PBS for 30min (adding 0.2% saponin for saponin-
permeabilized samples). Cells were incubated with primary and
secondary antibodies for 1hour at RT; antibodies were diluted in PBS
containing 1% goat serum (adding 0.2% saponin for saponin-
permeabilized samples) as detailed in the “Antibodies” section. Sam-
ples not meant for expansion microscopy were counterstained with
Table 1. Primary antibodies. IgG, immunoglobulin G.
Antibody Supplier Target Host
species Dilution
ab273073 Abcam S protein Human 1:400
NB100-56569 Novus
Biologicals M protein Rabbit 1:200
NBP2-41061 Novus
Biologicals E protein Rabbit 1:200
MA1-7403
Thermo
Fisher
Scientific
N protein Mouse
IgG2b 1:20
Ab01299-2.0 Absolute
Antibody dsRNA Mouse
IgG2a 1:200
ab22649 Abcam GM130 Rabbit 1:50
ab125006 Abcam LMAN-1 Rabbit 1:50
ab24170 Abcam LAMP1 Rabbit 1:100
ab22595 Abcam Calnexin Rabbit 1:200
ab131205 Abcam -Tubulin Mouse IgG1 1:200
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DAPI (4′,6-diamidino-2-phenylindole) (Abcam, ab228549) diluted
1:1000 in PBS for 15min at RT and mounted on Fisherbrand glass
microscope slides (Fisher Scientific, catalog no. 11572203) using the
VECTASHIELD Vibrance mounting reagent (2B Scientific).
Expansion microscopy
The fixed immunostained samples were expanded following a
published procedure (47) and imaged either on a confocal or on a
light sheet microscope as previously reported (48). Briefly, immuno-
stained cells were incubated with 0.25% glutaraldehyde in PBS
for 15 min, washed three times with PBS, and then incubated
with monomer solution (1× PBS, 2M NaCl, 2.5% acrylamide,
0.15% N,N′- methylenebisacrylamide, and 8.625% sodium acrylate)
for ~2min at RT. Gelation was started inverting coverslips onto
a drop of 150l of gelling solution [monomer solution/10%
N,N,N′,N′-tetramethylethylenediamine (TEMED)/10% ammonium
persulfate (APS), mixed in ratio 96:2:2] and left to gelate for 1hour
at RT in a humidified environment. Gels were digested in digestion
buffer [1× tris-acetate-EDTA buffer (TAE), 0.5% Triton X-100, and
20 mM CaCl2] containing proteinase K (~8 U/ml) overnight at 37°C.
Gels were eventually placed in double-distilled water to expand. The
expansion factor (4.2) was calculated as previously reported (48).
Microscopes
Wide-field microscope
Wide-field imaging of fixed SARS-CoV-2–infected cells was carried out
on a custom-built automated wide-field microscope. Frame (IX83,
Olympus), stage (Prior), Z drift compensator (IX3-ZDC2, Olympus),
four-wavelength high-power light-emitting diode light source
(LED4D067, Thorlabs), and camera (Zyla sCMOS, Andor) were con-
trolled by Micro-Manager (49). Respective filter cubes for DAPI (filter
set 49000-ET-DAPI, Chroma), Alexa Fluor 488 (filter set 49002-ET-
EGFP, Chroma), Alexa Fluor 568 (filter set 49008-ET-mCherry, Texas
Red, Chroma), Alexa Fluor 647 and Atto 647N (excitation filter
628/40, dichroic beam splitter Di02-R635, emission filter 708/75,
Semrock), as well as Atto 490LS (filter set 49003-ET-EYFP, Chroma;
emission filter replaced by 600LP, Semrock) were used. Images were
acquired with an Olympus U Plan Apo 60×/1.42 NA (numerical aper-
ture) oil objective lens at 30 to 35 random positions for each sample.
Confocal microscopes
The imaging of nonexpanded fixed samples was performed on a Zeiss
LSM 800 microscope using a Plan-Apochromat 63×/1.4 NA oil
objective. The microscope was controlled using the Zen software (ver-
sion 2.6), and, for acquisition of 16-bit images, a pinhole size of 1.0 Airy
unit (AU) for each channel, a scan speed of 5 (1.47 s per pixel), and
four times averaging were used. Pixel size was 70.6nm. Expanded gels
were cut to fit in a round glass-bottom dish (Ibidi -dish, catalog no.
81158) precoated with poly-l-lysine and were imaged on a Leica SP5
microscope using an apochromatic 63×/1.2 NA water objective. Im-
ages were acquired using a scanning frequency of 10Hz and a pixel size
ranging from 100 to 150nm. To increase the collection of signal from
the samples, the pinhole size was opened to 2.0 AU (in contrast to the
preset value of 1.0 AU), which corresponds to an optical section of 1.5 m.
Light sheet microscope
Expanded samples were imaged on a custom-built inverted selective
plane illumination microscope (iSPIM). Parts were purchased from
Applied Scientific Instrumentation (ASI) including the controller
(TG8_BASIC), scanner unit (MM-SCAN_1.2), right-angle objec-
tive mounting (SPIM-K2), stage (MS-2K-SPIM) with motorized Z
support (100-mm travel range; Dual-LS-100-FTP), and filter wheel
(FW-1000-8). All components were controlled by Micro-Manager
by means of the diSPIM plugin. The setup was equipped with a 0.3 NA
excitation objective (10×, 3.5-mm working distance; Nikon) and
a higher, 0.9 NA detection objective (W Plan-Apochromat 63×,
2.4mm working distance; Zeiss) to increase spatial resolution and
fluorescence signal collection. Lasers (OBIS445-75 LX, OBIS488-
150 LS, OBIS561-150 LS, and OBIS647-120 LX, Coherent) were
fiber-coupled into the scanner unit. An sCMOS camera (ORCA-Flash
4.0, Hamamatsu) was used to capture fluorescence. Respective
emission filters were BrightLineFF01-474/27, BrightLineFF01-540/50,
BrightLineFF01-609/54, and BrightLineFF0-708/75 (Semrock). Gels
containing expanded samples were cut into small strips and mounted
onto 24mm by 50mm rectangular coverslips with expanded cells
facing upward using Loctite super glue (Henkel), as previously re-
ported (48). The sample was then placed into an imaging chamber
(ASI, I-3078-2450), which was filled with double-distilled water. We
recorded volumes with a plane spacing of 0.5 m. Raw data were
deskewed using a custom MATLAB routine including a denoising
step to remove hot pixels. Stacks were automatically separated in the
respective color channels and were individually processed. Maximum
intensity projections were generated of the deskewed stacks.
Correlative structured illumination and atomic force microscope
Correlative AFM/fluorescence microscopy measurements were per-
formed as described before (50). AFM measurements were performed
Table 2. Secondary antibodies.
Antibody Supplier Target Conjugate Host species Dilution
A-11013 Thermo Fisher Scientific Human IgG Alexa Fluor 488 Goat 1:200
A-11011 Thermo Fisher Scientific Rabbit IgG Alexa Fluor 568 Goat 1:200
A-11031 Thermo Fisher Scientific Mouse IgG Alexa Fluor 568 Goat 1:100
A-21144 Thermo Fisher Scientific Mouse IgG2b Alexa Fluor 568 Goat 1:200
A-21244 Thermo Fisher Scientific Rabbit IgG Alexa Fluor 647 Goat 1:100
40839 Merck Rabbit IgG Atto 647N Goat 1:200
50185 Merck Mouse IgG Atto 647N Goat 1:200
610-156-040 Rockland Mouse IgG1 Atto 647N Goat 1:100
610-156-041 Rockland Mouse IgG2a Atto 647N Goat 1:100
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on a BioScope Resolve AFM (Bruker), operated in PeakForce Quanti-
tative Nanoscale Mechanical Characterization (QNM) mode, which
was combined with a custom-built structured illumination microscopy
system (51). A 60×/1.2 NA water immersion lens (UPLSAPO 60XW,
Olympus) was used for fluorescence excitation and detection, which
was captured with an sCMOS camera (C11440, Hamamatsu). The
wavelengths used for excitation were 488nm (iBEAM-SMART-488,
Toptica), 561nm (OBIS 561, Coherent), and 640nm [diode laser
module (MLD), Cobolt]. Images were acquired using customized
structure illumination microscopy (SIM) software.
Deconvolution
Confocal images and deskewed light sheet microscopy data of ex-
panded samples were deconvolved using the PSF Generator and
DeconvolutionLab2 plugins in Fiji (52). In total, 25 to 100 iterations
of the Richardson-Lucy algorithm were used. Deconvolved data
were maximum intensity projected in Fiji, optionally using color to
indicate depth.
Replication kinetics from wide-field data
The percentage of cells expressing each of the structural proteins of
SARS-CoV-2 and dsRNA was calculated semiautomatically using the
image-processing program Fiji (53). Cells expressing SARS-CoV-2
proteins and dsRNA were counted manually, whereas the total
number of cells was determined automatically using the “Analyze
particles” function. Images of the DAPI-stained nuclei were filtered
using the “Subtract background” (rolling ball radius=20 pixels),
“Gaussian blur” (sigma=15 pixels), and “Unsharp Mask” (radius=
10 pixels, mask weight=0.8) functions. Otsu thresholding was used
to create a binary mask image. Dividing cells and cells at the edges of
the image were excluded from analysis. For each time point, 30 to 35
wide-field microscope images (1000 to 1500 cells) were counted. For
the control time point 0 hpi, only around 250 cells were counted. To
determine the average expression levels of each of the structural pro-
teins of SARS-CoV-2, the infected cells were segmented manually
and the average fluorescence intensity in each viral protein channel
was measured. From each value, the mean background intensity was
subtracted and data were normalized using the highest average in-
tensity value of the respective time point (usually at 12 or 24 hpi) for
each protein. For each time point and protein, more than 80 cells
were analyzed, except the early time point of 5 hpi with only around
40 cells because the fraction of infected cells was very low (5 hpi:
n=46, 7.5 hpi: n=84, 10 hpi: n=85, 12 hpi: n=88, and 24 hpi:
n=81 for N, S, and M; and 5 hpi: n=40, 7.5 hpi: n=104, 10 hpi: n=83,
12 hpi: n=94, and 24 hpi: n=82 for E).
Stage kinetics from wide-field data
The OpenCV and scikit-image Python libraries were used for anal-
ysis. Quantification was performed on a dataset of ~35 wide-field
images per time point stained for the SARS-CoV-2N protein, the
SARS-CoV-2 S protein, GM130 (Golgi apparatus), and nucleus (DAP I).
Infection stages were assigned to each infected cell in the following
way. First, binary masks for the cell nuclei were created by using
local Otsu thresholding followed by contour detection and filtering
(see “Fragmentation analysis” section). For N protein detection, global
Otsu thresholding was applied to the corresponding channel. Then,
for each detected nucleus, the nucleus masks were used to create thin
perinuclear regions around the edge of each nucleus by upscaling
each mask by a factor of 1.15 and subtracting the original mask,
producing thin hoops around each nucleus. The cell was counted as
containing the N protein if more than one pixel in this region was
above the threshold value. The S protein analysis was the same, ex-
cept a fixed threshold value was used instead of Otsu thresholding,
and the masks were upscaled by a factor of 1.2 to produce a thicker
perinuclear region, as S protein signal was sparser than that of the
N protein. If cells were positive for N protein, but not S protein, they
were classified as stage 1. To distinguish between stages 2 and 3, a
fragmentation analysis of the Golgi apparatus (see “Fragmentation
analysis” section) was performed.
The accuracy of the method was checked by manually counting
the fraction of cells with the N protein and S protein signal as well as
the fraction with a fragmented Golgi apparatus at ~10 to 15 images
at 5 and 10 hpi. The results produced by the algorithm were within
1 SD of the mean values determined by manual analysis.
Image segmentation
The OpenCV and scikit-image Python libraries were used for the
segmentation. Wide-field and confocal images were segmented to
enable cell-specific analysis of the dataset as follows. Initially, all
channels of the images were merged to a grayscale image and the
background was removed via Li thresholding (54). Connected com-
ponent analysis was performed to segment the single-cell units in
the image. To be segmented as an object of interest, a connected
cluster was filtered via a minimum size of ~150 m2 (corresponds to
30,000 pixels for confocal images; the size of a typical cell nucleus
was ~200 to 250 m2). In case of high cell density or staining of ex-
tended structures (e.g., microtubules), connected component analy-
sis might lead to large numbers of cells being detected as one cluster.
Here, when a maximum cell cluster size of 1,000,000 pixels was ex-
tended, the number of nuclei in the cluster was isolated using the
DAPI-stained nuclei. For each nucleus in the cluster, the distances
to its K nearest neighboring nuclei were measured (usually use K=2
or K=3, given that in most cases <10 cells make up one cluster). The
cell outline of each single-cell unit was then defined by the outlines
of the nucleus (DAPI channel) and the half-distances to its K nearest
neighbors (choosing the maximum sized box that included all men-
tioned positions). For images characterized by low cell density, the
described methods successfully segmented all cells that can be iden-
tified manually. For high–cell density images or including extended
cell structures, these methods led to a good estimation of the cell
outline for the majority of cells (>75% by visual inspection).
Colocalization analysis
We quantified the spatial correlation between all four viral structural
proteins by measuring Spearman’s rank coefficients. The Spearman
coefficient is based on the ranking of image intensities. After assign-
ing ranks to the pixel intensity values in each of the two channels,
the Pearson correlation, which measures the degree of correlative
variation, between the rank values of the pixel intensities in the two
images is calculated. We also calculated the Manders coefficient
that, in contrast to the Spearman coefficient, measures co-occurrence
of intensities in the two channels rather than their correlation (55).
However, interpretation of the Manders coefficient can be difficult
because it depends on the ratio of total intensities in both channels.
In contrast to the Spearman coefficient, the Manders coefficient is
also affected by out-of-focus signal.
Spearman’s rank coefficients and Manders overlap coefficients were
computed by ColocAnalyzer. ColocAnalyzer is a custom program for
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Scherer et al., Sci. Adv. 8, eabl4895 (2022) 7 January 2022
SCIENCE ADVANCES | RESEARCH ARTICLE
14 of 17
image filtering and colocalization analysis, which is free and available he re:
https://github.com/LAG-MNG-CambridgeUniversity/ColocAnalyzer.
First, we saved images in such a manner that each of the channels of
interest fell into one of three main colors: red, green, or blue. Then,
we chose the two channels of interest (for example, red + blue or
green + red) to be analyzed. For each image, Otsu thresholding was
applied before computing colocalization coefficients on the remain-
ing pixels with higher intensities.
Spearman’s rank coefficients were computed by ColocAnalyzer as
= 1 − 6 ∑
q d
q
2
─
n( n 2 − 1)
Here, dq = rank [I1(q)] − rank [I2(q)]is the difference between
ranks computed for pixel q in channel 1 and in channel 2 independently.
n is the number of pixels that were analyzed. Because, after thresh-
olding, a substantial fraction of pixels was blanked (would have zero
intensity), we used only those pixels that had nonzero values in both
channels to avoid an impact from black pixels.
The Manders overlap coefficient was computed by ColocAnalyzer
using the formula provided in the original paper (56)
MOC = ∑
q
Np I1(q ) * I2(q)
──────────────
√
___________________
∑
q
Np I1 (q) 2 * ∑
q I2 (q) 2
where I1(q) and I2(q) are the intensities of pixel q in the first and
second channel, respectively. Np is the total number of pixels taken
for analysis.
Spot detection and analysis
N protein, dsRNA, and lysosome spot detection and analysis from
microscopy images were performed by a customized MATLAB routine.
For spot detection, we first applied median filtering to the image:
Each pixel intensity value is decreased by a median value of intensi-
ties in a subarea of 60 pixels by 60 pixels around this pixel (the size
of this subarea was chosen empirically). After Otsu thresholding of
the filtered image, we determined the positions of the connected
pixels with nonzero intensities. We called each cluster of such con-
nected pixels a spot. Last, we filtered out spots that were smaller
than 300nm in diameter (approximately corresponds to Abbe’s res-
olution limit) and had a mean intensity value smaller than 10% of
the maximum intensity. The area of each detected spot was calculated
from the number of pixels per spot (the pixel size was 107.3 nm).
For spot shape analysis, we fitted each spot to an ellipse with the
customized MATLAB routine “fit_ellipse.m” (www.mathworks.com/
matlabcentral/fileexchange/3215-fit_ellipse) and used the two radii
Rmin and Rmax obtained from fitting to compute the eccentricity value of
each spot: e =
√
_
1 −
(
R min
_
R max
)
2 , where Rmin and Rmax are the smaller and
larger radii of the ellipse, respectively. The distance between N pro-
tein and dsRNA spots was calculated as the minimal distance be-
tween the two spot centers.
The spot-to-spot distance between S protein and the lysosome
marker LAMP1 was analyzed using the spot colocalization plugin
ComDet for Fiji. For particle detection within the plugin, particle
sizes between 3 and 4 pixels (corresponds to 210 to 280 nm) and an
intensity threshold of 3 to 10 SDs of the average particle intensity
were selected. The maximum distance between colocalized spots
was set to 4 pixels (corresponding to 280 nm).
Fragmentation analysis
Fragmentation analysis of the Golgi apparatus was performed on
16-bit wide-field images. The OpenCV and scikit-image Python
libraries were used for the analysis. First, the channel with the DAPI-
stained nuclei was segmented into cell nuclei and background
using local Otsu thresholding followed by contour detection using
the cv2.findContours function within the OpenCV library. The
detected contours were filtered by size and circularity to ensure
only the single nonoverlapping nuclei were selected. Specifically,
contours with lengths in the range of 250 to 2500 pixels (30 to 300 m)
were accepted. From manual inspection, the nuclei contours fell
roughly within the range of 300 to 700 pixels (35 to 85 m). Only
contours with length-to-area ratios of less than 0.05 were se-
lected to eliminate non-elliptical shapes. The contours were then
scaled down to 90% of their original size to avoid overlap with
structures from other channels and filled to produce a mask for
each image.
Next, local Otsu thresholding was performed on the Golgi appa-
ratus channel. The mask of the corresponding nucleus was sub-
tracted from the result. A rectangular region was created around
each detected nucleus for subsequent location of the Golgi appara-
tus. The size of the region was determined by first creating a
rectangle such that its borders were tangential to the outline of the
detected nucleus and then scaling up its size by a factor of 2. Con-
tour detection was performed within each region to locate the Golgi
apparatus or its fragments. The angular size of each contour with
respect to the center of the corresponding nucleus was calculated. It
was found that a fragmented Golgi apparatus was typically detected
as a single contour because thresholding of the wide-field images
did not resolve the large number of small fragments, so only the size
of the largest detected fragment for each corresponding nucleus was
recorded. Contours with the angular size of less than 20° were found
to be indistinguishable from noise, and so, the corresponding cells
were excluded from the analysis.
Microtubule directionality analysis
Directionality of microtubules was computed by a custom MATLAB
routine on the basis of the texture detection technique introduced in
(57). The method relies on computing gray-level co-occurrence
matrices (GLCMs) as proposed in (58). The matrix is defined for
single values of pixel position shifts [dx, dy] and consists of relative
frequencies pij that two pixels with gray levels i and j are separated
by [dx, dy]. For eight-bit images, the GLCM will be a matrix of 256 ×
256 elements. Instead of using [dx, dy], we used the concept of angle
and distance: [φ, d]. We varied the distances from 10 to 100 pixels in
5-pixel steps (19 values). The minimum of 10 pixels corresponds to
approximately 1 m, so that short microtubules less than 1 m in
length were excluded from the analysis. The range of directions φ =
[0°:180°] was divided into 45 segments with 4° steps for fine resolu-
tion of directionality. In total, we generated 19 × 45=855 GLCMs
for each image. Then, as in (57), we computed the joint probability
of occurrence for the specified pixel pair
T(φ, d ) = ∑
i
∑
j
p ij (i − μ x ) (i − μ y )
───────────
σ x σ y
where x, y, x, and y are the means and SDs of px and py, respectively:
p x = ∑
i p ij , p y = ∑
j p ij . Next, we averaged those values across distanc-
es to leave only the angular dependence
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Scherer et al., Sci. Adv. 8, eabl4895 (2022) 7 January 2022
SCIENCE ADVANCES | RESEARCH ARTICLE
15 of 17
T(φ ) =
∑
d T(φ, d)
─
19
Then, we obtained the texture correlation values H(φ) by nor-
malizing the joint probability for each direction
H(φ ) = T(φ)
─
∑
i=0
45 T(4i)
The texture correlation function shows greater values for the angles
with preferable directions in microtubule images. Visual inspection
on a number of microtubule images showed good performance of
the method and its ability to find precisely (up to 4° in our case)
dominating microtubule directions in the image. Last, the direc-
tionality coefficient was computed from summing up the second
moments around each peak, from valley to valley
D = 1 − ∑
p
n p
∑
φϵ w p [ (φ − φ p ) 2 H(φ)]
where np is the number of peaks in H(φ), φp is the value of an angle
at the pth peak, wp is the range for the pth peak between two valleys,
and is the normalizing coefficient =
1
_
45 1
____________
∑
p
n p ∑
φϵ w p (φ − φ p ) 2
.
Cell stiffness measurement and analysis
For AFM cell stiffness measurements, Vero cells were plated at 60%
confluence in 50-mm glass-bottom dishes (GWST-5040, WillCo
Wells BV) the day before infection, infected, and fixed as described
before. Live cell probes (PFQNM-LC, Bruker AFM probes) were used
for all experiments. The probes were precalibrated for spring con-
stant (nominal of 0.08 N/m), and deflection sensitivity was calibrated
at the start of each experiment. The force applied to the cells was kept
constant throughout the experiments, with typical values ranging
between 150 and 300 pN. Force curves were fitted to a Hertz model
F =
4
√
_
R c
─
3 E
─
1 − 2
3/2
where Rc is the radius of tip curvature, v is the sample’s Poisson’s
ratio, E is the Young’s modulus, and is the indentation depth. Curve
fitting and Young’s modulus calculation were performed using
nanoscope analysis.
Data visualization and statistical analysis
Graphs were plotted with GraphPad Prism. Statistical significance
between two values was determined using a two-tailed, unpaired
Student’s t test (GraphPad Prism). In the figures, asterisks denote
statistical significance as calculated by Student’s t test (*P<0.05,
**P<0.01, ***P<0.001, and ****P<0.0001).
SUPPLEMENTARY MATERIALS
Supplementary material for this article is available at https://science.org/doi/10.1126/
sciadv.abl4895
View/request a protocol for this paper from Bio-protocol.
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Scherer et al., Sci. Adv. 8, eabl4895 (2022) 7 January 2022
SCIENCE ADVANCES | RESEARCH ARTICLE
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Acknowledgments: We thank R. Ulferts for help in selecting the SARS-CoV-2 antibodies and
J. D. Manton for help with the alignment of the light sheet microscope. Funding: C.F.K.
acknowledges funding from EPSRC (EP/L015889/1 and EP/H018301/1), Wellcome Trust
(3-3249/Z/16/Z and 089703/Z/09/Z), MRC (MR/K015850/1 and MR/K02292X/1), AstraZeneca,
and Infinitus (China) Ltd. J.L.H. receives funding from NIHR/UKRI. The SARS-CoV-2 work
performed at the Laboratory of Viral Zoonotics (LVY), University of Cambridge, was carried out
with support from the UKRI/DHSC grant COV0170-HICC: Humoral Immune Correlates for
COVID19: Defining protective responses and critical readouts for clinical trials (G107217),
awarded to J.L.H. and the LVZ. Author contributions: C.F.K., K.M.S., and L.M. conceptualized
the project. G.W.C. prepared samples for all experimental data. K.M.S., L.M., L.C.S.W., and A.F.-V.
stained all samples and performed all imaging experiments. G.W.C., H.S., M.S.S., and C.L.G.
performed plaque assays and PCR experiments. K.M.S., L.M., S.M., M.Br., and M.Ba. performed
data analysis. I.M. performed and evaluated AFM experiments. J.R.L. provided code for light
sheet data processing. K.M.S., L.M., G.W.C., L.C.S.W., S.M., M.Br., M.Ba., I.M., H.S., and C.F.K.
contributed to manuscript writing. All authors revised the article. Competing interests: The
authors declare that they have no competing interests. Data and materials availability: The
BetaCoV/Australia/VIC01/2020 strain of SARS-CoV-2 can be provided by the Victorian
Infectious Diseases Reference Laboratory, Melbourne, through Public Health England’s
pending scientific review and a completed material transfer agreement. Requests for a sample
of the SARS-CoV-2 strain should be submitted to Public Health England. The code for the
ColocAnalyzer can be downloaded from Apollo, the University of Cambridge Repository, at
https://doi.org/10.17863/CAM.77726. Original data that were analyzed with the code are
available at https://doi.org/10.17863/CAM.78118. All data needed to evaluate the conclusions
in the paper are present in the paper and/or the Supplementary Materials.
Submitted 19 July 2021
Accepted 15 November 2021
Published 7 January 2022
10.1126/sciadv.abl4895
Downloaded from https://www.science.org on February 14, 2022
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SARS-CoV-2 nucleocapsid protein adheres to replication organelles before viral
assembly at the Golgi/ERGIC and lysosome-mediated egress
Katharina M. SchererLuca MascheroniGeorge W. CarnellLucia C. S. WunderlichStanislaw MakarchukMarius
BrockhoffIoanna MelaAna Fernandez-VillegasMax BarysevichHazel StewartMaria Suau SansCharlotte L. GeorgeJacob R.
LambGabriele S. Kaminski-SchierleJonathan L. HeeneyClemens F. Kaminski
Sci. Adv., 8 (1), eabl4895. • DOI: 10.1126/sciadv.abl4895
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