Study details how Covid causes blood clots
https://doi.org/10.1042/BSR20210611
Received: 12 March 2021
Revised: 13 July 2021
Accepted: 29 July 2021
Accepted Manuscript Online:
30 July 2021
Version of Record published:
20 August 2021
Research Article
SARS-CoV-2 spike protein S1 induces fibrin(ogen)
resistant to fibrinolysis: implications for microclot
formation in COVID-19
Lize M. Grobbelaar1, Chantelle Venter1, Mare Vlok2, Malebogo Ngoepe3,4, Gert Jacobus Laubscher5,
Petrus Johannes Lourens5, Janami Steenkamp1,6, Douglas B. Kell1,7,8 and Etheresia Pretorius1
1Department of Physiological Sciences, Faculty of Science, Stellenbosch University, Stellenbosch, Private Bag X1, Matieland 7602, South Africa; 2Central Analytical Facility: Mass
Spectrometry Stellenbosch University, Tygerberg Campus, Room 6054, Clinical Building, Francie van Zijl Drive, Tygerberg, Cape Town 7505, South Africa; 3Department of
Mechanical Engineering, Faculty of Engineering and the Built Environment, University of Cape Town, Cape Town, Rondebosch 7701, South Africa; 4Stellenbosch Institute for
Advanced Study, Wallenberg Research Centre, Stellenbosch University, Stellenbosch, South Africa; 5Private practice clinician, Mediclinic Stellenbosch, Stellenbosch 7600, South
Africa; 6PathCare Laboratories, PathCare Business Centre, PathCare Park, Neels Bothma Street, N1 City 7460, South Africa; 7Department of Biochemistry and Systems Biology,
Institute of Systems, Molecular and Integrative Biology, Faculty of Health and Life Sciences, University of Liverpool, Liverpool L69 7ZB, U.K.; 8The Novo Nordisk Foundation Centre
for Biosustainability, Technical University of Denmark, Kemitorvet 200, Kgs Lyngby 2800, Denmark
Correspondence: Etheresia Pretorius (resiap@sun.ac.za) or Douglas B. Kell (dbk@liv.ac.uk)
Severe acute respiratory syndrome coronavirus 2 (SARS-Cov-2)-induced infection, the
cause of coronavirus disease 2019 (COVID-19), is characterized by unprecedented clinical
pathologies. One of the most important pathologies, is hypercoagulation and microclots
in the lungs of patients. Here we study the effect of isolated SARS-CoV-2 spike protein S1
subunit as potential inflammagen sui generis. Using scanning electron and fluorescence microscopy
as well as mass spectrometry, we investigate the potential of this inflammagen
to interact with platelets and fibrin(ogen) directly to cause blood hypercoagulation. Using
platelet-poor plasma (PPP), we show that spike protein may interfere with blood flow. Mass
spectrometry also showed that when spike protein S1 is added to healthy PPP, it results
in structural changes to β and γ fibrin(ogen), complement 3, and prothrombin. These proteins
were substantially resistant to trypsinization, in the presence of spike protein S1. Here
we suggest that, in part, the presence of spike protein in circulation may contribute to the
hypercoagulation in COVID-19 positive patients and may cause substantial impairment of
fibrinolysis. Such lytic impairment may result in the persistent largemicroclotswe have noted
here and previously in plasma samples of COVID-19 patients. This observation may have
important clinical relevance in the treatment of hypercoagulability in COVID-19 patients.
Introduction
Severe acute respiratory syndrome coronavirus 2 (SARS-Cov-2)-induced infection, the cause of coronavirus
disease 2019 (COVID-19), is characterized by unprecedented clinical pathologies. Phenotypic vascular
characteristics are strongly associated with various coagulopathies that may result in either bleeding
and thrombocytopenia or hypercoagulation and thrombosis [1,2]. Various circulating and dysregulated
inflammatory coagulation biomarkers, including fibrin(ogen), D-dimer, P-selectin and von Willebrand
Factor (VWF), C-reactive protein (CRP), and various cytokines, directly bind to endothelial receptors.
Endotheliopathies are therefore a key clinical feature of the condition [3,4]. During the progression of the
various stages of the COVID-19, markers of viral replication, as well as VWF and fibrinogen depletion
with increased D-dimer levels and dysregulated P-selectin levels, followed by a cytokine storm, are likely
to be indicative of a poor prognosis [5–8]. This poor prognosis is further worsened as together with a
©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution
License 4.0 (CC BY).
1
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
Figure 1. Schematic representation of SARS-CoV-2 Spike glycoprotein
Adapted from[21]. Abbreviations: HR1, heptad repeat 1; HR2, heptad repeat 2; S1, subunit 1; S2, subunit 2. This image was created
with BioRender (https://biorender.com/).
substantial deposition of microclots in the lungs [9–11]. Plasma of COVID-19 patients also carries a massive load of
preformed amyloid clots [5], and there are also numerous reports of damage to erythrocytes [12–14], platelets, and
dysregulation of inflammatory biomarkers [5–8].
The virulenceof thepathogen is closelylinkedtoitsmembraneproteins.One suchprotein, foundontheCOVID-19
virus, is the spike protein, which is a membrane glycoprotein. The spike proteins are the key factors for virus attachment
to target cells, as they bind to the angiotensin-converting 2 (ACE2) surface receptors [15]. Spike proteins are
class I viral fusion proteins [16]. They are present as protruding homotrimers on the viral surface and mediate virus
entry into the target host cells [17]. A singular spike protein is between 180 and 200 kDa in size and contains an extracellularN-
terminal, a transmembrane domain fixed in themembrane of the virus, and a short intracellularC-terminal
segment [16,18]. Spike proteins are coated with polysaccharide molecules that serve as camouflage. This helps evade
surveillance by the host immune system during entry [18]. The S1 subunit is responsible for receptor binding [19],
with subunit 2 (S2), a carboxyl-terminal subunit, responsible for viral fusion and entry [20] (see Figure 1).
Receptor binding is certainly responsible for cell-mediated pathologies, but does not itself explain the coagulopathies.
Spike protein, can however be shed, and it has been detected in various organs, including the urinary tract
[22]. S1 proteins can also cross the blood–brain barrier [23]. Free S1 particles may also play a role in the pathogenesis
of the disease [24,25]. Free spike protein can potentially be released due to spontaneous ‘firing’ of the S protein trimers
on the surface of virions, and infected cells liberates free receptor binding domain-containing S1 particles [24].
Here we study the effect of isolated SARS-CoV-2 spike protein S1 subunit as potential pro-inflammatory inflammagen
sui generis.We investigate the potential of this inflammagen to directly interactwith platelets and fibrin(ogen)
to cause fibrin(ogen) protein changes and blood hypercoagulation.We also determine if the spike protein may interfere
with blood flow, by comparing na¨ıve healthy platelet-poor plasma (PPP) samples, with and without added spike
protein, to PPP samples from COVID-19 positive patients (before treatment). We conclude that the spike protein
may have pathological effects directly, without being taken up by cells. This provides further evidence that targeting
it directly, whether via vaccines or antibodies, is likely to be of therapeutic benefit.
2 ©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution
License 4.0 (CC BY).
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
Materials and methods
Sample demographics and considerations
Blood was collected from healthy volunteers (n=11; 3 males and 8 females; mean age: 43.4 +−
11.7 years) to serve
as controls. Individuals who smoke, who were diagnosed with cardiovascular diseases, clotting disorders (coagulopathies),
and/or any known inflammatory conditions (e.g. Type 2 DiabetesMellitus (T2DM), rheumatoid arthritis,
tuberculosis, asthma, etc.) could not serve as control volunteers. Furthermore, pregnancy, lactation, hormonal therapy,
oral contraceptive usage, and/or using anticoagulants, were also factors that resulted in exclusion. Smoking was
excluded since it has been proven to impair coagulation, fibrinolysis, and the hemostatic process [26]. Microfluidics
analysis included a preliminary analysis using PPP samples from two COVID-19 positive patients, on the day of first
diagnosis and before any treatment was given. Both patients were diagnosed with moderate to severe COVID-19
symptoms (1 male and 1 female, mean age: 78.5+−
7.7 years).
Blood sample collection
Either a qualified phlebotomist or medical practitioner drew the volunteers’ (control) blood via venepuncture, adhering
to standard sterile protocol. Blood samples were stored in two to three 4.5-ml sodium citrate (3.2%) tubes
(BD Vacutainer®, 369714). After several gentle inversions, the collected citrate tubes were allowed to rest at room
temperature foraminimumof 30min to allowfor adequate anticoagulant amalgamation before commencing sample
preparation.Whole blood (WB) was centrifuged at 3000×g for 15 min at room temperature to isolate erythrocytes.
The supernatant, i.e. PPP, was collected and stored in 1.5-ml Eppendorf tubes at −80◦C, till needed for experiments.
Spike protein preparation
SARS-CoV-2 (2019-nCoV) Spike protein S1 Subunit, mFcTag, was purchased from Sino Biological (Beijing, China)
(catalog number 40591-V05H1) and prepared using double distilled water, following the instructions provided. A
total of 400 μl diluent (endotoxin-free water) was added to the 100 μg spike protein to create a stock solution (A) of
0.25 mg.ml−1. This stock solution was diluted to working solutions. To determine the concentration of spike protein
needed to result in significant, but yet a high enough concentration to cause physiological effects on the viscoelastic
properties of blood, different concentrations of spike protein in PPP were assessed with fluorescence microscopy. A
healthy control blood sample were separated into four 1.5-ml Eppendorf tubes with different final exposure concentrations
of spike protein in the PPP of 100, 50, 10, and 1 ng.ml−1. PPP samples were incubated with the various spike
protein concentrations for 30 min at room temperature.
Fluorescence microscopy of purified fibrinogen and PPP with and without
thrombin
Concentration verification
To verify which spike protein concentration will be effective, 5 μl of the PPP exposed to the varies spike protein
concentrations was placed on a glass slide, after being exposed to the fluorescent amyloid dye, Thioflavin T (ThT)
(Sigma–Aldrich, St. Louis,MO,U.S.A.) for 30min at roomtemperature. The final concentration of ThT in all prepared
sampleswas 0.005mM.After the evaluation of the samples, with the varying spike protein concentrations, itwas found
that the final exposure concentration of 1 ng.ml−1 was sufficient and used for the rest of the study.
Amyloid protein and anomalous clotting in PPP samples
To study spontaneous anomalous clotting of fibrin(ogen), in the na¨ıve healthy PPP samples, and in the presence of
spike protein, 5 μl PPP exposed to 1 ng.ml−1 (final concentrations) of spike protein, was smeared on a glass slide and
covered with a coverslip. This was done after it was exposed to the fluorescent amyloid dye, ThT (final concentration:
0.005mM)(Sigma–Aldrich, St. Louis,MO,U.S.A.) for 30 min at roomtemperature. Fibrin PPP clots, with andwithout
spike protein and after exposure to ThT, were also prepared by adding 2.5 μl of thrombin (7 U.ml−1, South African
National Blood Service) to 5 μl PPP and was placed on a glass slide and covered with a coverslip. The excitation
wavelength for ThT was set at 450–488 nm and the emission at 499–529 nm and processed samples were viewed
using a Zeiss Axio Observer 7 fluorescentmicroscope with a Plan-Apochromat 63×/1.4 Oil DIC M27 objective (Carl
Zeiss Microscopy,Munich, Germany).
Fluorescence microscopy images of healthy PPP with and without spike protein were analyzed using Fiji® (Java
1.6 0 24 [64-bit]) to numerically represent the images. The total area of fluorescing particles or anomalous clotting
(identified by the amyloid dye, ThT) [27–29] was determined using a thresholding algorithm in Fiji®. Images were
firstly set to scale in Fiji® according to the magnification of the lens used on the fluorescent microscope, followed
©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution
License 4.0 (CC BY).
3
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
by the selection of appropriate threshold value to regard as much of the foreground and disregard as much of the
background of the image as possible. In order to optimize the amount of images thresholded, a program was written
in Java to simultaneously analyze a group of images (see SupplementaryMaterial). The total percentage of anomalous
clots in each image was calculated and the average of all the images per sample was calculated. These average values
were used for statistical analysis.
Purified fibrin(ogen) clot model
To determine if spike protein causes changes in purified fibrinogen, our purified fibrin(ogen) clotmodel of choice was
fluorescent fibrinogen conjugated to Alexa Fluor™ 488 (Thermo Fisher, F13191). A final fibrinogen concentration of
2 mg.ml−1 was prepared in endotoxin-free water and exposed to 1 ng.ml−1 (final concentrations) spike protein for
30 min at room temperature. A total of 5 μl of the purified fibrinogen was placed on a glass slide, with 2.5 μl of
thrombin. The excitation wavelength for our fluorescent fibrinogen model was set at 450–488 nm and the emission
at 499–529 nm and processed samples were viewed using a Zeiss Axio Observer 7 fluorescent microscope with a
Plan-Apochromat 63×/1.4 Oil DIC M27 objective (Carl Zeiss Microscopy, Munich, Germany).
WB (hematocrit)
Hematocrit was exposed to a final exposure concentration of 1 ng.ml−1 spike protein. The fluorescentmarker, CD62P
(platelet surface P-selectin) was added to the hematocrit to study platelet activation. CD62P is found on the membrane
of platelets and then translocated to the platelet membrane surface. The translocation occurs after the platelet
P-selectin is released fromthe cellular granules during platelet activation [5,7]. A total of 4 μl CD62P (PE-conjugated)
(IM1759U, Beckman Coulter, Brea, CA, U.S.A.) was added to 20 μl WB (either na¨ıve or incubated with spike protein).
The hematocrit exposed to the marker was incubated for 30 min (protected from light) at room temperature.
The excitation wavelength for the CD62P was 540–570 nm and the emission 577–607 nm. Processed samples were
also viewed using a Zeiss Axio Observer 7 fluorescent microscope with a Plan-Apochromat 63×/1.4 Oil DIC M27
objective (Carl Zeiss Microscopy, Munich, Germany).
Scanning electron microscopy of WB samples
Scanning electron microscopy (SEM) was used to view healthyWB samples, with and without the addition of spike
protein. A total of 10 μl WB was placed on a glass coverslip and prepared according to previously published SEM
preparation methods [30,31], starting with washing steps in phosphate-buffered saline (PBS) (pH = 7.4) (Thermo
Fisher Scientific, 11594516) for 20 min. Fixation was performed by coating the slides in 4% formaldehyde (FA) for
30 min, followed by washing them in PBS three times. For each wash, the PBS should be left on for 3 min before
removing and washing again. Osmiumtetroxide (OsO4) (Sigma–Aldrich, 75632) was added for 15 min and the slides
were washed in PBS three-times with 3 min in each once more. The next step was to serially dehydrate the slides
with ethanol followed by a drying step using hexamethyldisilazane (HMDS) (Sigma–Aldrich, 379212). Samples were
mounted on glass slides and coatedwith carbon.The slides were viewed on a ZeissMERLINFE-SEMwith the InLens
detector at 1 kV (Carl Zeiss Microscopy, Munich, Germany).
Microfluidics
Microfluidic analysis was performed using healthy PPP and healthy pooled PPP samples (three pooled PPP samples),
with and without spike protein, and two COVID-19 PPP samples. Pooled samples were used due to the volume
required for this experiment.
Hardware
Amicrofluidic setup was used to simulate and investigate clot growth under conditions of flow. A Cellix microfluidic
syringe pump (Cellix Ltd, Dublin, Ireland) was used to drive flow through Cellix Vena8 Fluoro+ biochips (Cellix Ltd,
Dublin, Ireland), comprising eight straight microfluidic flow channels each, at flow rates specified in the following
paragraph.Asinglemicrochannel had awidth of 400 μmandaheight of 100 μm(equivalent diameter of 207 μm), and
a length of 2.8 cm. The dimensions of themicrochannel were in the same order of magnitude as those of some vessels
of themicrovasculature, that is, under 300 μm [32]. In order to observe clot evolution in real-time, the biochips were
placed under the Zeiss Axio Observer 7 fluorescent microscope with a 10×/0.25 objective (Carl Zeiss Microscopy,
Munich, Germany).
4 ©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution
License 4.0 (CC BY).
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
Figure 2. Experimental protocol for growing clots in a microfluidics system
Flow conditions
A new flow channel was used for every run. The channel was flushed with distilled water at a flow rate of 1 ml.min−1
for 1 min. After 5 min, thrombin was infused through the microchannel at a flow rate of 50 μl.min−1 for 90 s (Figure
2). The channel was left to stand for another 5 min and then the sample (control, control with spike, or COVID-19)
was infused at a flow rate of 10 μl.min−1 for 5 min, with a video recording of the channel (Figure 2). The flow was
then stopped after 5min, and a set of micrographs was taken across the channel. The sample was then left for another
5min, to see if any additional changes occur (Figure 2). This flow rate corresponds with a shear rate, γ ≈ 250 s−1 and
a Reynolds number, Re ≤ 1. One of the main challenges in attempting to achieve consistent shear rates and Reynolds
number was the variability in viscosity from sample to sample. Furthermore, blood flowing through microvessels is
known to behave in a non-Newtonianmanner, adding to the complexities of variable viscosity within a single sample.
To achieve standardization between samples, a constant flow rate was used.
Proteomics
Four healthy PPP samples were analyzed before and after addition of spike protein. The samples were diluted in 10
mMammoniumbicarbonate to 1mg.ml−1. A total of 1 μg trypsin (NewEngland Biosystems) was added to the plasma
for 1:50 enzyme to substrate ratio. No reduction or alkylation was performed.
Liquid chromatography
Dionex nano-RSLC
Liquid chromatography was performed on a Thermo ScientificUltimate 3000 RSLC equipped with a 5mm×300 μm
C18 trap column (Thermo Scientific) and a CSH 25 cm × 75 μm, 1.7 μmparticle size C18 column (Waters) analytical
column. The solvent systememployed was loading: 2% acetonitrile:water; 0.1% FA; Solvent A: 2% acetonitrile: water;
0.1% FA and Solvent B: 100% acetonitrile:water. The samples were loaded on to the trap column using loading solvent
at a flow rate of 2 μl/min from a temperature controlled autosampler set at 7◦C. Loading was performed for 5 min
before the sample was eluted onto the analytical column. Flow rate was set to 300 nl/min and the gradient generated
©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution
License 4.0 (CC BY).
5
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
as follows: 5.0–30% B over 60 min and 30–50% B from 60–80 min. Chromatography was performed at 45◦C and the
outflow delivered to the mass spectrometer.
Mass spectrometry
Mass spectrometry was performed using a Thermo Scientific Fusion mass spectrometer equipped with a Nanospray
Flex ionization source. Plasma samples, before and after addition of spike protein addition (1 ng.ml−1 final exposure
concentration), four of our control samples were analyzed with this method. The sample was introduced through a
stainless-steel nano-bore emitter.Datawere collected inpositivemodewith spray voltage set to1.8kVandion transfer
capillary set to 275◦C. Spectra were internally calibrated using polysiloxane ions at m/z =445.12003. MS1 scans were
performed using the orbitrap detector set at 120000 resolution over the scan range 375–1500 with AGC target at 4 E5
and maximum injection time of 50 ms. Data were acquired in profile mode. MS2 acquisitions were performed using
monoisotopic precursor selection for ion with charges +2 to +7 with error tolerance set to+ −10 ppm. Precursor ions
were excluded from fragmentation once for a period of 60 s. Precursor ions were selected for fragmentation in HCD
mode using the quadrupole mass analyzer with HCD energy set to 30%. Fragment ions were detected in the Orbitrap
mass analyzer set to 30000 resolution. The AGC target was set to 5E4 and the maximum injection time to 100 ms.
The data were acquired in centroid mode.
Statistical analysis
Data analysis: plasma samples
Statistical analyses of data generated were performed using GraphPad Prism software (version 9.0.0). The normality
of the data was assessed using the Shapiro–Wilk normality test. For analysis of data between two groups, paired t tests
(for pairwise statistical comparisons between data from untreated and treated control groups) and unpaired t tests (for
non-pairwise statistical comparisons) were performed to assess statistical significance for parametric data, whereas
the Mann–Whitney test was utilized to test for statistical significance in non-parametric data and the Wilcoxon’s
test was used for significance in paired parametric data. When comparing three or more experimental groups, the
Kruskal–Wallis test (non-parametric data) or one-wayANOVA(parametric data) test was applied to test for statistical
significance.AP-value of less than 0.05 was considered to be statistically significant. Parametric datawere presented as
the mean and standard deviation (SD), whereas non-parametric data were presented as the median and interquartile
range (IQR).
Mass spectrometer data analysis
The raw files generated by the mass spectrometer were imported into Proteome Discoverer v1.4 (Thermo Scientific)
and processed using the Sequest and Amanda algorithms. Database interrogation was performed against the
2019-nCOVpFASTA1 database. Semi-tryptic cleavage with two missed cleavages was allowed for. Precursor mass
tolerance was set to 10 ppm and fragment mass tolerance set to 0.02 Da. Demamidation (NQ), oxidation (M) were
allowed as dynamic modifications. Peptide validation was performed using the Target-Decoy PSM validator node.
The search results were imported into Scaffold Q+ for further validation (www.proteomesoftware.com) andstatistical
testing. A t test was performed on the datasets and the emPAI quantitative method used to compare the datasets.
Supplementary material and raw data
All supplementary material and raw data can be accessed here: https://1drv.ms/u/s!
AgoCOmY3bkKHisg5J0nb6wqsBzzWAQ?e=lmObMy
Results
Fluorescence microscopy of purified fibrinogen and PPP
Fluorescence microscopy was utilized to visualize the fluorescent amyloid signals in spontaneously formed anomalous
clots, present in a fluorescent fibrinogen model, and also in healthy PPP with and without spike protein. As a
preliminary investigation PPP from a single healthy control were incubated for 30 min with varying spike protein
concentrations, followed by 30-min incubation with ThT and lastly preparation for viewing. It was found that the
final exposure concentrations of 1 ng.ml−1 was sufficient and used for the rest of the study (see supplementary raw
data for other exposure concentrations’ micrographs).
Figure 3A,B shows representative micrographs of purified fluorescent (Alexa Fluor™488) fibrinogen with added
thrombin and after exposure to 1 ng.ml−1 spike protein. A denser fibrin clot formed in the presence of spike protein
(Figure 3B). In PPP, with and without thrombin, the green fluorescent ThT signal indicates areas of amyloid deposit
6 ©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution
License 4.0 (CC BY).
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
Figure 3. Representative fluorescence micrographs of purified fluorescent (Alexa Fluor™ 88) fibrinogen (note no ThT added)
with added thrombin to form extensive fibrin clots
(A) Fluorescent fibrinogen with thrombin; (B) fluorescent fibrinogen with added spike protein (final exposure concentration 1 ng.ml−1)
and thrombin.
Table 1 Percentage average amyloid area in PPP with and without spike protein and with and without thrombin
Na¨ıve healthy PPP vs na¨ıve healthy PPP + added thrombin (n=10)
P-value (Wilcoxon’s test, paired non-parametric data expressed as median [Q1 − Q3]0.2
Median of na¨ıve healthy samples 0.3% [0.1–0.8%]
Median of na¨ıve control samples (+ thrombin) 0.9% [0.3–1.5%]
Na¨ıve healthy PPP + spike protein (1 ng.ml−1) vs healthy PPP + spike protein (1 ng.ml−1) + added thrombin) (n=10)
P-value (data normally distributed; paired t test) 0.3
Mean percentage amyloid of healthy samples + spike 1.9% (+−
1.2%)
Mean percentage amyloid of healthy samples + spike (+ Thrombin) 2.4% (+−
1.3%)
Na¨ıve healthy PPP vs healthy PPP + spike protein (1 ng.ml−1) (n=10)
P-value (Wilcoxon’s test, paired non-parametric data expressed as median [Q1 − Q3]0.004 (*)
Median of healthy samples 0.3% [0.1–0.8%]
Median of healthy samples + spike 1.9% [1.2–2.4%]
Na¨ıve healthy PPP + added thrombin vs healthy PPP + spike protein (1 ng.ml−1) + added thrombin) (n=10)
P-value (data normally distributed; paired t test) 0.0036 (*)
Mean percentage amyloid of control samples (+ Thrombin) 0.9% (+−
0.6%)
Mean percentage amyloid of control samples + spike (+ Thrombin) 2.4% (1.3%)
Statistical significance was established at P<0.05. (* = P<0.01). Data are represented as either mean +−
SD or median [Q1 − Q3].
formation. ThT is known to bind to open hydrophobic areas in protein and these are amyloid in nature [27,33–35].
Figure 4A,D shows representative micrographs of a healthy PPP smear, with and without thrombin, and with added
ThT where slight anomalous clotting is seen. In contrast, when spike protein is added to PPP, with and without thrombin,
amajor increase in dense anomalous clotted deposits, with an amyloid nature, were noted (referred to as amyloid
deposits) (Figure 4B,D). Athresholding algorithmwas applied to themicrographs (with and without thrombin) using
Fiji® which was used to calculate the total area of amyloid deposits in each micrograph (in total, 320 micrographs
were analyzed). Using this method, the average total percentage of amyloid deposits per group was calculated (na¨ıve
healthy PPP, na¨ıve healthy PPP + thrombin, and PPP incubated with 1 ng.ml−1 spike protein, with and without added
thrombin) (see Table 1).As expected, therewere no significant differences between%area amyloid deposits of healthy
PPP with and without thrombin However, there was a significant increase in % area amyloid deposits in PPP before
and after added spike protein, in both PPP smears and fibrin clots (where thrombin was added).
©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution
License 4.0 (CC BY).
7
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
Figure 4. Representative fluorescence micrographs of PPP from healthy individuals after addition of ThT (green fluorescent
signal)
(A) PPP smear. (B) PPP with spike protein. (C) PPP with thrombin to create extensive fibrin clot. (D) PPP exposed to spike protein
followed by addition of thrombin. Final spike protein concentration was 1 ng.ml−1.
Platelet activity
Fluorescence microscopy was used to visualize platelet activation in naive healthy hematocrit and hematocrit incubatedwith
spike protein (1 ng.ml−1 final concentration). Sampleswere incubated with the plateletmarker, CD62P-PE.
Figure 5A shows representative platelets from na¨ıve control samples, while Figure 5B shows micrographs after spike
protein incubation. Spike protein caused an increase in platelet hyperactivation (Figure 5B arrows).
SEM of WB
SEMwas used to assess the erythrocyte and platelet ultrastructure after treatment with spike protein (1 ng.ml−1 final
concentration). Figure 6A,B shows micrographs fromhealthyWBsamples, while Figure 6C–H shows micrographs of
8 ©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution
License 4.0 (CC BY).
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
Figure 5. Fluorescence microscopy micrographs of platelets, before and after exposure to spike protein
(A) Representative platelets from hematocrit incubated with fluorescent marker, CD62P-PE. (B) Representative micrographs showing
activated platelets after exposure to spike protein. The white arrows point to hyperactivated activated platelets. White arrows
show hyperactivated platelets clumping together.
WB after incubation with spike protein. The majority of erythrocytes from healthy untreated controls were normocytic
(regularly shaped) [Figure 6A (arrow)], and featured characteristic discoid shapes smooth, regular membrane
surfaces. Slight platelet activation is seen due to contact activation (Figure 6B). TheWB incubated with spike protein
showed erythrocyte agglutination, despite the very low concentration of the spike protein. An increase in platelet hyperactivation,
membrane spreading (Figure 6C,D), platelet-derived microparticle formation were noted due to spike
protein exposure. The formation of spontaneous and anomalous fibrin(ogen) deposits with an amyloid nature, were
prominent in all the samples incubated with spike protein, without the addition of thrombin [Figure 6E–H (arrows)].
Microfluidics
Figure 7 show the clots that formed in the flow chambers after 5 min of starting the experiment. Healthy PPP formed
a small clot along the bottom surface of the channel, as seen in Figure 7A. In healthy plasma, clot formation was a
relatively slow and gradual process, resulting in the formation of a modest clot (see Supplementary healthy PPP video
S1). Clots formed in healthy PPP were relatively small and were limited to the walls of the flow channel. The clot had
orderly layers that did not disrupt flow through the centre of the channel. As expected, clot formation was also less
frequent than with the other samples (Figure 7B).The PPP with added spike protein showed a combination of a fibrous
laminar clot and disorderly clottedmass (Figure 7E,F) (see Supplementary healthy PPP with added spike protein video
S2). The COVID-19 PPP show disorderly clots that cover the bulk of the channel and often protruding into the center
of the flow channel and disrupting flow (Figure 7C,D). In COVID-19 PPP, the reaction between thrombin and PPP
occurred rapidly, resulting in large clots after approximately 90 s (see Supplementary pooled COVID-19 patient PPP
video S3). Interestingly, these clots did not propagatemuch after the initial burst, indicating thatmost of the thrombin
was consumed in a short period of time. Clots also formed with the PPP with the addition of the spike protein, but
not as disruptive as the COVID-19 PPP clots.
An interesting observation was that clots from healthy PPP could easily be dislodged by flushing the flow channel
with water at a rate of 1 ml.min−1 (0.42 m.s−1). Similarly, clots from healthy PPP with added spike protein could be
dislodged in a similar fashion. COVID-19 clots, on the contrary, could not be displaced or dislodged and remained
intact, even with the force of high-speed water flow in a small flow channel. This observation was consistent for all
the COVID-19 samples.
Mass spectrometry analysis
Figure 8 shows results from the mass spectrometry analysis. Mass spectrometry showed that when spike protein
is added to healthy PPP, it results in structural changes to β and γ fibrin(ogen), complement 3, and prothrombin.
©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution
License 4.0 (CC BY).
9
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
Figure 6. Whole blood sample of healthy volunteers, before and after exposure to spike protein
(A–H) Representative scanning electron micrographs of healthy control WB, with and without spike protein. (A,B) Healthy WB
smears, with arrow indicating normal erythrocyte ultrastructure. (C–H) Healthy WB exposed to spike protein (1 ng.ml−1 final concentration),
with (C,D) indicating the activated platelets (arrow), (E,F) showing the spontaneously formed fibrin network and (G,H)
the anomalous deposits that is amyloid in nature (arrows) (scale bars: (E) 20 μm; (A) 10 μm; (F,G) 5 μm; (H) 2 μm; (C) 1 μm; (B,D)
500 nm).
10 ©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution
License 4.0 (CC BY).
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
Figure 7. Representative micrographs of PPP clots in the microfluidic chambers (black horizontal lines are the outlines of
the chambers) that were coated with thrombin
(A) Healthy PPP clot, with small clot formation (arrow), with (B) no clot formed in the healthy PPP sample. (C,D) examples of clots
from COVID-19 PPP samples and (E,F) healthy PPP clot with spike protein. Black arrow = small clot formed in control sample; red
arrows large clots in COVID-19 sample.
©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution
License 4.0 (CC BY).
11
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
Figure 8. Mass spectrometry showing PPP of four samples with and without added spike protein
Spike protein results in structural changes to β and γ fibrin(ogen), complement 3, and prothrombin.
These proteins were substantially resistant to trypsinization, in the presence of spike protein (for sequence data, see
supplementary files).
Discussion
In this laboratory analysis, we provide evidence that spike protein does indeed play amajor role in hypercoagulability
seen in COVID-19 patients. It causes anomalous clotting in both purified fluorescent fibrinogen and in PPP from
healthy individuals, where the nature of the clots were shown to be amyloid (ThT as our amyloid dye of choice). An
interesting observation was that these dense deposits were noted both in smears exposed to spike protein, and when
thrombin was added. The addition of thrombin causes purified (Alexa Fluor™ 488) fibrinogen to polymerize into
fibrin networks. Typically, these networks are net-like (Figure 3A). In the presence of spike protein, the structure
changed to form dense clot deposits (Figure 3B). These deposits were seen in our fluorescent fibrin(ogen) model
and PPP from healthy individuals exposed to spike protein. In healthy PPP exposed to spike protein, followed by
incubation with ThT, there was a significant increase in anomalous clots with an amyloid nature, (Figure 4D), when
compared with the healthy PPP. In the current paper, we did not analyze blood samples and clotting propensity of
PPP fromother patient cohorts, e.g. those with bacterial pneumonia or other acute viral diseases.However, our group
(and others) have previously studied blood from HIV patients, where significant hypercoagulation in this patient
cohort was noted [36–39]. We have also, recently, reported significant hypercoagulation in patients suffering from
long COVID/PASC [40].
Spike protein also caused major ultrastructural changes in WB (as viewed with the SEM), where platelet hyperactivation
was noted (Figure 6C,D). Increased in spontaneously formed fibrin network, as well as anomalous clot
formation were also observed in SEM micrographs (Figure 6E–H). Interestingly, extensive spontaneous fibrin network
formation was noted, without the addition of thrombin. This is in line with results that were recently published,
where we showed similar ultrastructure in blood smears form COVID-19 positive patients. In these patients, platelet
hyperactivation, anomalous clotting with amyloid signal, and spontaneous fibrin fiber formation were also observed
[6,7].
With the microfluidics flow system, clots were formed, by infusing the entire microchannel with thrombin, thus
simulating a hypercoagulable state, where endothelial damage was extensive. Given that the flow channel was made
entirely of plastic and was devoid of any endothelial cells, the main component under investigation was the PPP
(mostly fibrinogen protein) itself, which, in the case of the COVID-19 samples, may have contained downstream effects
of some endothelial changes that would give rise to the hypercoagulable state that is characteristic of the disease.
The flow setup used in the present study could not directly account for endothelial changes but nonetheless demonstrated
that COVID-19 also results in changes in the clotting profile of the PPP. This was evident in the rapid rate of
thrombin consumption and fibrin formation in COVID-19 clots, and also in the nature of the PPP clots that were
formed.
The clots thatwere observed in the healthy PPP with added spike protein,were of particular interest as they demonstrated
a bridge between healthy PPP clots and COVID-19 clots. As described in the ‘Results’ section, the healthy PPP
clots were relatively small and orderly, while COVID-19 PPP clots were large, disorderly masses that formed rapidly
12 ©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution
License 4.0 (CC BY).
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
Figure 9. Simplified coagulation diagram depicting healthy and pathological processes
(1A) The intrinsic and (1B) extrinsic pathways converge into the (1C) common pathway. These pathways lead to the conversion of
soluble fibrinogen to insoluble fibrin, catalyzed by thrombin. (2) Tissue plasminogen activator (tPA) or urokinase-type plasminogen
activator (uPA) converts plasminogen into plasmin. A healthy fibrinolytic system regulates the coagulation pathway and assists with
successful lysis of the insoluble fibrin clot. (3) Plasmin cleaves fibrin into fibrin degradation products (FDPs), including D-dimer. (4)
Protein C and thrombomodulin both regulate coagulation: thrombin binds to its receptor, thrombomodulin, resulting in activated
protein C (APC). APC then inhibits both Va and VIIIa. (5) Dysregulated inflammatory molecules may interfere with tissue factor (TF)
expression. (6) Dysregulated inflammatory molecules may also down-regulate thrombomodulin, resulting in hypercoagulation, as
Va and VIIIa activities are then not sufficiently modulated. (7) In our laboraoty suty, we added Spike protein S1 to healthy plasma.
Pathophysiology was noted in both prothrombin and fibrinogen chains. (8) Dysregulated inflammatory molecules in circulation can
inhibit of the fibrinolytic system via up-regulation of plasminogen activator inhibitor-1 (PAI-1). PAI-I up-regulation interferes with tPA
function, and ultimately results in a dysregulated coagulation system. (9) α2AP inhibits plasmin and ultimately will prevent sufficient
fibrinolysis to happen. (Figure created with Biorender.com).
©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution
License 4.0 (CC BY).
13
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
anddisruptedPPPflowinthe channel. ThehealthyPPPclotswith added spikeprotein,were a combinationof the two,
demonstrating disorderly clumped clot areas, coexisting with laminar fibrous PPP clots (which were larger than the
healthy PPP clots). This intermediate state may arise froma number of factors, including the interaction of other biological
actors which were absent from the flow setup and the time of exposure to spike protein. Further investigations
would be beneficial for understanding the clotting mechanisms that are altered in the presence of spike protein.
One of the obvious differences, which was inadvertently observed while trying to clean the channels with
high-speed water flow (i.e. by mechanical means), was the ease of healthy PPP and healthy PPP with added spike
protein clot dissolution. However, there was a complete failure to dislodge or disturb COVID-19 PPP clots from the
channels. Given the clot lysis and dissolution is a complex interplay between biochemical and biophysical factors,
investigation of the effects of different therapeutic agents could elucidate this phenomenon [41]. The flow protocol
used in this study would be a useful platformfor testing different treatments for clinical application. A further limitation
of this exploration is the use of PPP in investigating clot formation at a scale appropriate to themicrovasculature.
While the protocol enables the study of fibrin microclots, which are of interest in COVID-19, it excludes the influence
of RBCs, which are known to heavily influence the non-Newtonian flow behavior of blood at that scale [42]. The
inaccuracy of the flow regime arising from this exclusion and from the variability of viscosity introduces error into
the results. Nonetheless, the inclusion of flow an appropriate spatial scale has enabled us to observe COVID-19 PPP
clot formation over space and time, under dynamic conditions, and has given insights which would otherwise prove
difficult to glean.
A further avenue for exploration would include examining clot stability and removal for other patients with acute
inflammatory responses arising from acute infections. Longstaff and co-workers found increased fibrinolytic resistance
in inflammatory conditions arising from acute infection [43]. Examining this phenomenon in ourmicrofluidic
setup would be beneficial.
Given thatmicroclots can blockmicrocapillaries and thereby inhibit oxygen exchange,we have recently also studied
plasma samples, using proteomics results from Long COVID/PASC, T2DM, with acute COVID-19 and compared
resultswith plasma samples fromhealthy individuals. Interestingly, plasma fromT2DMand formhealthy individuals,
immediately digested fully after a first trypsinization step, however, persistent microclots remained in the plasma
samples from Long COVID/PASC and from acute COVID-19 samples, still contained large anomalous (amyloid)
deposits (microclots) [40]. After a second trypsinization, the persistent pellet deposits were solubilized.We detected
various inflammatorymolecules that are substantially increased in both the supernatant and trapped in the solubilized
pellet deposits of acute COVID-19 and Long COVID/PASC, versus the equivalent volume of fully digested fluid of the
control samples and T2DM[40].Of particular interest was a substantial increase in α(2)-antiplasmin (α2AP), various
fibrinogen chains in both acute COVID-19 and Long COVID/PASC digested microclots. A comparison between
healthy plasma and acute COVID-19 solubilized clots also showed a significant increase in coagulation factor XIII A
chain, VWF Complement component C7 and CRP [40].
In the current study, mass spectrometry confirmed that spike protein causes structural changes to β and γ fibrin(
ogen), complement 3, and prothrombin. These proteins become less resistant to trypsinization and changes the
conformation, in such a way that there is a significant difference in peptide structure before and after spike protein
addition. The current results therefore confirm results we have found in our recent proteomics analysis [40].
Our recent data suggest that there is an increase in (2)-antiplasmin inside the microclots of both acute COVID-19
and Long COVID, and we also note pathophysiology in the fibrinogen chains. In the present paper, we could induce
fibrinogen chain pathology, after adding spike protein to healthy plasma. To result in an increase in molecules like
α2-antiplasmin and VWF (and others), in patients with acute COVID-19 and also those with Long COVID/PASC,
many physiological pathways should be activated. See Figure 9 (adjusted from[40, 44–46]) for a visual representation
of the coagulation pathway and where it may be affected by S1.
Conclusion
Scanning electron- and fluorescence microscopy revealed large, dense anomalous and amyloid masses in WB and
PPP of healthy individuals where spike protein was added to the samples. Mass spectrometry confirmed that when
spike protein was added to PPP, it interacts with plasma proteins, resulting in fibrin(ogen), prothrombin and other
proteins linked to coagulation, to become substantially resistant to trypsinization, resulting in less fragments. Flow
analysis confirmed that microclots may impair blood flow. Here we suggest that, in part, the presence of spike protein
S1 in circulation may contribute to the hypercoagulation in COVID-19 positive patients and may cause severe
impairment of fibrinolysis. The effects of S2might also be relevant, and should also be investigated in future. It is also
now accepted that coagulation pathology is central in acute COVID-19 [47–52]. Several autopsy results have also
14 ©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution
License 4.0 (CC BY).
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
confirmed microthrombi throughout the lung and associated with right ventricular dilation of the heart. Recently,
Ackermann and co-workers reported that histologic analysis of pulmonary vessels in patients with COVID-19 shows
widespread thrombosis withmicroangiopathy [4]. The authors also showed that alveolar capillarymicrothrombiwere
nine-times as prevalent in patients with COVID-19 as in patients with influenza (P<0.001). Fibrinolytic impairment
may therefore be the direct cause of the largemicroclots we have noted here in SEMand fluorescence microscopy,
and previously in plasma samples of COVID-19 patients [6,7]. Clotting pathologies in acute COVID-19 infection
might therefore benefit from following a regime of continued anticlotting therapy to support the fibrinolytic system
function [40].
Data Availability
All supplementary material and raw data can be accessed here: https://1drv.ms/u/s!AgoCOmY3bkKHisg5J0nb6wqsBzzWAQ?e=
XAsc7w
Competing Interests
The authors declare that there are no competing interests associated with the manuscript.
Funding
This work was supported by the Novo Nordisk Foundation [grant number NNF10CC1016517 (to Douglas B. Kell)].
Open Access
Open access for this article was enabled by the participation of University of Liverpool in an all-inclusive Read & Publish pilot with
Portland Press and the Biochemical Society under a transformative agreement with JISC.
CRediT Author Contribution
Lize M. Grobbelaar: Formal analysis, Visualization, Writing—original draft. Chantelle Venter: Data curation, Project administration.
Mare Vlok: Software, Formal analysis, Investigation. Malebogo Ngoepe: Methodology. Gert Jacobus Laubscher:
Writing—review and editing. Petrus Johannes Lourens: Writing—review and editing. Janami Steenkamp: Project administration,
Writing—review and editing. Douglas B Kell: Writing—review and editing. Etheresia Pretorius: Conceptualization, Resources,
Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Project administration,
Writing—review and editing.
Ethics Approval
Ethical clearance for the study was obtained from the Health Research Ethics Committee (HREC) of Stellenbosch University
(South Africa) (reference: N19/03/043, project ID: 9521). The experimental objectives, risks, and details were explained to volunteers
both verbally and in text and informed consent were obtained prior to blood collection. Strict compliance to ethical guidelines
and principles Declaration of Helsinki, South African Guidelines for Good Clinical Practice, and Medical Research Council
Ethical Guidelines for Research were kept for the duration of the study and for all research protocols.
Consent for Publication
All authors approved submission of the paper.
Abbreviations
COVID-19, coronavirus disease 2019; CRP, C-reactive protein; FA, formaldehyde; PBS, phosphate-buffered saline; PPP,
platelet-poor plasma; SARS-CoV-2, severe acute respiratory syndrome coronavirus 2; SEM, scanning electron microscopy; ThT,
Thioflavin T; T2DM, Type 2 diabetes mellitus; VWF, von Willebrand Factor; WB, whole blood.
References
1 Gupta, A., Madhavan, M.V., Sehgal, K., Nair, N., Mahajan, S., Sehrawat, T.S. et al. (2020) Extrapulmonary manifestations of COVID-19. Nat. Med. 26,
1017–1032, https://doi.org/10.1038/s41591-020-0968-3
2 Perico, L., Benigni, A., Casiraghi, F., Ng, L.F.P., Renia, L. and Remuzzi, G. (2021) Immunity, endothelial injury and complement-induced coagulopathy in
COVID-19. Nat. Rev. Nephrol. 17, 46–64, https://doi.org/10.1038/s41581-020-00357-4
3 Goshua, G., Pine, A.B., Meizlish, M.L., Chang, C.H., Zhang, H., Bahel, P. et al. (2020) Endotheliopathy in COVID-19-associated coagulopathy: evidence
from a single-centre, cross-sectional study. Lancet Haematol. e575–e582, https://doi.org/10.1016/S2352-3026(20)30216-7
4 Ackermann, M., Verleden, S.E., Kuehnel, M., Haverich, A., Welte, T., Laenger, F. et al. (2020) Pulmonary vascular endothelialitis, thrombosis, and
angiogenesis in Covid-19. N. Engl. J. Med. 383, 120–128, https://doi.org/10.1056/NEJMoa2015432
©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons
Attribution License 4.0 (CC BY).
15
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
5 Grobler, C., Maphumulo, S.C., Grobbelaar, L.M., Bredenkamp, J.C., Laubscher, G.J., Lourens, P.J. et al. (2020) Covid-19: The rollercoaster of
fibrin(ogen), D-dimer, von Willebrand Factor, P-selectin and their interactions with endothelial cells, platelets and erythrocytes. Int. J. Mol. Sci. 21,
5168, https://doi.org/10.3390/ijms21145168
6 Pretorius, E., Venter, C., Laubscher, G.J., Lourens, P.J., Steenkamp, J. and Kell, D.B. (2020) Prevalence of readily detected amyloid blood clots in
‘unclotted’ Type 2 diabetes mellitus and COVID-19 plasma: a preliminary report. Cardiovasc. Diabetol. 19, 193,
https://doi.org/10.1186/s12933-020-01165-7
7 Venter, C., Bezuidenhout, J.A., Laubscher, G.J., Lourens, P.J., Steenkamp, J., Kell, D.B. et al. (2020) Erythrocyte, platelet, serum ferritin, and P-selectin
pathophysiology implicated in severe hypercoagulation and vascular complications in COVID-19. Int. J. Mol. Sci. 21, 8234,
https://doi.org/10.3390/ijms21218234
8 Roberts, I., Muelas, M.W., Taylor, J.M., Davison, A.S., Xu, Y., Grixti, J.M. et al. (2020) Untargeted metabolomics of COVID-19 patient serum reveals
potential prognostic markers of both severity and outcome. medRxiv, https://doi.org/10.1101/2020.12.09.20246389
9 Renzi, S., Landoni, G., Zangrillo, A. and Ciceri, F. (2020) MicroCLOTS pathophysiology in COVID 19. Korean J. Intern. Med.,
https://doi.org/10.3904/kjim.2020.336
10 Ciceri, F., Beretta, L., Scandroglio, A.M., Colombo, S., Landoni, G., Ruggeri, A. et al. (2020) Microvascular COVID-19 lung vessels obstructive
thromboinflammatory syndrome (MicroCLOTS): an atypical acute respiratory distress syndrome working hypothesis. Crit. Care Resusc. 22, 95–97
11 Bobrova, L., Kozlovskaya, N., Korotchaeva, Y., Bobkova, I., Kamyshova, E. and Moiseev, S. (2020) Microvascular COVID-19 lung vessels obstructive
thromboinflammatory syndrome (MicroCLOTS): a new variant of thrombotic microangiopathy? Crit. Care Resusc. 22, 284
12 Lam, L.M., Murphy, S.J., Kuri-Cervantes, L., Weisman, A.R., Ittner, C.A.G., Reilly, J.P. et al. (2020) Erythrocytes reveal complement activation in patients
with COVID-19. medRxiv, https://doi.org/10.1101/2020.05.20.20104398
13 Berzuini, A., Bianco, C., Paccapelo, C., Bertolini, F., Gregato, G., Cattaneo, A. et al. (2020) Red cell-bound antibodies and transfusion requirements in
hospitalized patients with COVID-19. Blood 136, 766–768, https://doi.org/10.1182/blood.2020006695
14 Akhter, N., Ahmad, S., Alzahrani, F.A., Dar, S.A., Wahid, M., Haque, S. et al. (2020) Impact of COVID-19 on the cerebrovascular system and the
prevention of RBC lysis. Eur. Rev. Med. Pharmacol. Sci. 24, 10267–10278
15 Bergmann, C.C. and Silverman, R.H. (2020) COVID-19: coronavirus replication, pathogenesis, and therapeutic strategies. Cleve. Clin. J. Med. 87,
321–327, https://doi.org/10.3949/ccjm.87a.20047
16 Kawase, M., Kataoka, M., Shirato, K. and Matsuyama, S. (2019) Biochemical analysis of coronavirus spike glycoprotein conformational intermediates
during membrane fusion. J. Virol. 93, e00785–19, https://doi.org/10.1128/JVI.00785-19
17 Walls, A.C., Park, Y.J., Tortorici, M.A., Wall, A., McGuire, A.T. and Veesler, D. (2020) Structure, function, and antigenicity of the SARS-CoV-2 spike
glycoprotein. Cell 181, 281.e286–292.e286, https://doi.org/10.1016/j.cell.2020.02.058
18 Zhang, C., Zheng, W., Huang, X., Bell, E.W., Zhou, X. and Zhang, Y. (2020) Protein structure and sequence re-analysis of 2019-nCoV genome does not
indicate snakes as its intermediate host or the unique similarity between its spike protein insertions and HIV-1. bioRxiv 19, 1351–1360,
https://doi.org/10.1101/2020.02.04.933135
19 Watanabe, Y., Allen, J.D., Wrapp, D., McLellan, J.S. and Crispin, M. (2020) Site-specific glycan analysis of the SARS-CoV-2 spike. Science 369,
330–333, https://doi.org/10.1126/science.abb9983
20 Flores-Alanis, A., Sandner-Miranda, L., Delgado, G., Cravioto, A. and Morales-Espinosa, R. (2020) The receptor binding domain of SARS-CoV-2 spike
protein is the result of an ancestral recombination between the bat-CoV RaTG13 and the pangolin-CoV MP789. BMC Res. Notes 13, 398,
https://doi.org/10.1186/s13104-020-05242-8
21 Duan, L., Zheng, Q., Zhang, H., Niu, Y., Lou, Y. and Wang, H. (2020) The SARS-CoV-2 spike glycoprotein biosynthesis, structure, function, and
antigenicity: implications for the design of spike-based vaccine immunogens. Front. Immunol. 11, 576622,
https://doi.org/10.3389/fimmu.2020.576622
22 George, S., Pal, A.C., Gagnon, J., Timalsina, S., Singh, P., Vydyam, P. et al. (2021) Evidence for SARS-CoV-2 spike protein in the urine of COVID-19
patients. medRxiv, https://doi.org/10.1101/2021.01.27.21250637
23 Rhea, E.M., Logsdon, A.F., Hansen, K.M., Williams, L.M., Reed, M.J., Baumann, K.K. et al. (2021) The S1 protein of SARS-CoV-2 crosses the
blood–brain barrier in mice. Nat. Neurosci. 24, 368–378, https://doi.org/10.1038/s41593-020-00771-8
24 Letarov, A.V., Babenko, V.V. and Kulikov, E.E. (2020) Free SARS-CoV-2 spike protein S1 particles may play a role in the pathogenesis of COVID-19
infection. Biochemistry (Moscow) 86, 257–261, https://doi.org/10.1134/S0006297921030032
25 Buzhdygan, T.P., DeOre, B.J., Baldwin-Leclair, A., McGary, H., Razmpour, R., Galie, P.A. et al. (2020) The SARS-CoV-2 spike protein alters barrier
function in 2D static and 3D microfluidic in vitro models of the human blood–brain barrier. bioRxiv
26 Pretorius, E., Oberholzer, H.M., van der Spuy, W.J. and Meiring, J.H. (2010) Smoking and coagulation: the sticky fibrin phenomenon. Ultrastruct. Pathol.
34, 236–239, https://doi.org/10.3109/01913121003743716
27 Kell, D.B. and Pretorius, E. (2017) Proteins behaving badly. Substoichiometric molecular control and amplification of the initiation and nature of amyloid
fibril formation: lessons from and for blood clotting. Prog. Biophys. Mol. Biol. 123, 16–41, https://doi.org/10.1016/j.pbiomolbio.2016.08.006
28 Pretorius, E., Bester, J., Mbotwa, S., Robinson, C. and Kell, D.B. (2016) Acute induction of anomalous blood clotting by molecular amplification of highly
substoichiometric levels of bacterial lipopolysaccharide. J. R. Soc. Interface 13, 20160539, PMC5046953, https://doi.org/10.1098/rsif.2016.0539
29 Pretorius, E., Vermeulen, N., Bester, J., Lipinski, B. and Kell, D.B. (2013) A novel method for assessing the role of iron and its functional chelation in
fibrin fibril formation: the use of scanning electron microscopy. Toxicol. Mech. Methods 23, 352–359, https://doi.org/10.3109/15376516.2012.762082
30 Pretorius, E. (2013) The adaptability of red blood cells. Cardiovasc. Diabetol. 12, 63, https://doi.org/10.1186/1475-2840-12-63
31 Pretorius, E., Vermeulen, N., Bester, J. and Lipinski, B. (2013) Novel use of scanning electron microscopy for detection of iron-induced morphological
changes in human blood. Microsc. Res. Tech. 76, 268–271, https://doi.org/10.1002/jemt.22163
16 ©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons
Attribution License 4.0 (CC BY).
Bioscience Reports (2021) 41 BSR20210611
https://doi.org/10.1042/BSR20210611
32 Jacob, M., Chappell, D. and Becker, B.F. (2016) Regulation of blood flow and volume exchange across the microcirculation. Crit. Care 20, 319,
https://doi.org/10.1186/s13054-016-1485-0
33 Adams, B., Nunes, J.M., Page, M.J., Roberts, T., Carr, J., Nell, T.A. et al. (2019) Parkinson’s disease: a systemic inflammatory disease accompanied by
bacterial inflammagens. Front. Aging Neurosci. 11, 210, https://doi.org/10.3389/fnagi.2019.00210
34 de Waal, G.M., Engelbrecht, L., Davis, T., de Villiers, W.J.S., Kell, D.B. and Pretorius, E. (2018) Correlative light-electron microscopy detects
lipopolysaccharide and its association with fibrin fibres in Parkinson’s disease, Alzheimer’s disease and Type 2 diabetes mellitus. Sci. Rep. 8, 16798,
https://doi.org/10.1038/s41598-018-35009-y
35 Page, M.J., Thomson, G.J.A., Nunes, J.M., Engelbrecht, A.M., Nell, T.A., de Villiers, W.J.S. et al. (2019) Serum amyloid A binds to fibrin(ogen), promoting
fibrin amyloid formation. Sci. Rep. 9, 3102, https://doi.org/10.1038/s41598-019-39056-x
36 Jackson, B.S., Nunes Goncalves, J. and Pretorius, E. (2020) Comparison of pathological clotting using haematological, functional and morphological
investigations in HIV-positive and HIV-negative patients with deep vein thrombosis. Retrovirology 17, 14,
https://doi.org/10.1186/s12977-020-00523-3
37 Jackson, B.S. and Pretorius, E. (2019) Pathological clotting and deep vein thrombosis in patients with HIV. Semin. Thromb. Hemost. 45, 132–140,
https://doi.org/10.1055/s-0038-1676374
38 Teer, E., Joseph, D.E., Driescher, N., Nell, T.A., Dominick, L., Midgley, N. et al. (2019) HIV and cardiovascular diseases risk: exploring the interplay
between T-cell activation, coagulation, monocyte subsets, and lipid subclass alterations. Am. J. Physiol. Heart Circ. Physiol. 316, H1146–H1157,
https://doi.org/10.1152/ajpheart.00797.2018
39 Green, S.A., Smith, M., Hasley, R.B., Stephany, D., Harned, A., Nagashima, K. et al. (2015) Activated platelet-T-cell conjugates in peripheral blood of
patients with HIV infection: coupling coagulation/inflammation and T cells. AIDS 29, 1297–1308, https://doi.org/10.1097/QAD.0000000000000701
40 Pretorius, E., Vlok, M., Venter, C., Bezuidenhout, J.A., Laubscher, G.J., Steenkamp, J. et al. (2021) Persistent clotting protein pathology in Long
COVID/post-acute sequelae of COVID-19 (PASC) is accompanied by increased levels of antiplasmin. medRxiv,
https://doi.org/10.1101/2021.05.21.21257578
41 Hudson, N.E. (2017) Biophysical mechanisms mediating fibrin fiber lysis. Biomed Res. Int. 2017, 2748340, https://doi.org/10.1155/2017/2748340
42 McHedlishvili, G. (1998) Disturbed blood flow structuring as critical factor of hemorheological disorders in microcirculation. Clin. Hemorheol. Microcirc.
19, 315–325
43 Longstaff, C., Varju´ , I., So´ tonyi, P., Szabo´ , L., Krumrey, M., Hoell, A. et al. (2013) Mechanical stability and fibrinolytic resistance of clots containing fibrin,
DNA, and histones. J. Biol. Chem. 288, 6946–6956, https://doi.org/10.1074/jbc.M112.404301
44 Kell, D.B. and Pretorius, E. (2015) The simultaneous occurrence of both hypercoagulability and hypofibrinolysis in blood and serum during systemic
inflammation, and the roles of iron and fibrin(ogen). Integr. Biol. (Camb.) 7, 24–52, https://doi.org/10.1039/c4ib00173g
45 de Waal, G.M., de Villiers, W.J. and Pretorius, E. (2021) The link between bacterial inflammagens, leaky gut syndrome and colorectal cancer. Curr. Med.
Chem., https://doi.org/10.2174/0929867328666210219142737
46 Miszta, A., Huskens, D., Donkervoort, D., Roberts, M.J.M., Wolberg, A.S. and de Laat, B. (2021) Assessing plasmin generation in health and disease.
Int. J. Mol. Sci. 22, 2758, https://doi.org/10.3390/ijms22052758
47 Giannis, D., Ziogas, I.A. and Gianni, P. (2020) Coagulation disorders in coronavirus infected patients: COVID-19, SARS-CoV-1, MERS-CoV and lessons
from the past. J. Clin. Virol. 127, 104362, https://doi.org/10.1016/j.jcv.2020.104362
48 Kollias, A., Kyriakoulis, K.G., Dimakakos, E., Poulakou, G., Stergiou, G.S. and Syrigos, K. (2020) Thromboembolic risk and anticoagulant therapy in
COVID-19 patients: emerging evidence and call for action. Br. J. Haematol. 189, 846–847, https://doi.org/10.1111/bjh.16727
49 Middeldorp, S., Coppens, M., van Haaps, T.F., Foppen, M., Vlaar, A.P., Mu¨ ller, M.C.A. et al. (2020) Incidence of venous thromboembolism in hospitalized
patients with COVID-19. J. Thromb. Haemost. 18, 1995–2002, https://doi.org/10.1111/jth.14888
50 Miesbach, W. and Makris, M. (2020) COVID-19: coagulopathy, risk of thrombosis, and the rationale for anticoagulation. Clin. Appl. Thromb. Hemost. 26,
1076029620938149, https://doi.org/10.1177/1076029620938149
51 Levi, M., Thachil, J., Iba, T. and Levy, J.H. (2020) Coagulation abnormalities and thrombosis in patients with COVID-19. Lancet Haematol. 7,
e438–e440, https://doi.org/10.1016/S2352-3026(20)30145-9
52 Liu, P.P., Blet, A., Smyth, D. and Li, H. (2020) The science underlying COVID-19: implications for the cardiovascular system. Circulation 142, 68–78,
https://doi.org/10.1161/CIRCULATIONAHA.120.047549
©2021 The Author(s). This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution
License 4.0 (CC BY).
17