Mitigating the non-specific uptake of immunomagnetic microparticles enables the extraction of endothelium from human fat

Abstract

Endothelial cells are among the fundamental building blocks for vascular tissue engineering. However, a clinically viable source of endothelium has continued to elude the field. Here, we demonstrate the feasibility of sourcing autologous endothelium from human fat – an abundant and uniquely dispensable tissue that can be readily harvested with minimally invasive procedures. We investigate the challenges underlying the overgrowth of human adipose tissue-derived microvascular endothelial cells by stromal cells to facilitate the development of a reliable method for their acquisition. Magnet-assisted cell sorting strategies are established to mitigate the non-specific uptake of immunomagnetic microparticles, enabling the enrichment of endothelial cells to purities that prevent their overgrowth by stromal cells. This work delineates a reliable method for acquiring human adipose tissue-derived microvascular endothelial cells in large quantities with high purities that can be readily applied in future vascular tissue engineering applications.

Full text

ARTICLE
Mitigating the non-specific uptake of
immunomagnetic microparticles enables the
extraction of endothelium from human fat
Jeremy A. Antonyshyn 1,2, Vienna Mazzoli 1,2, Meghan J. McFadden1,2, Anthony O. Gramolini 2,3,
Stefan O. P. Hofer4,5, Craig A. Simmons1,2,6 & J. Paul Santerre 1,2,7 ✉
Endothelial cells are among the fundamental building blocks for vascular tissue engineering.
However, a clinically viable source of endothelium has continued to elude the field. Here, we
demonstrate the feasibility of sourcing autologous endothelium from human fat –an abun-
dant and uniquely dispensable tissue that can be readily harvested with minimally invasive
procedures. We investigate the challenges underlying the overgrowth of human adipose
tissue-derived microvascular endothelial cells by stromal cells to facilitate the development of
a reliable method for their acquisition. Magnet-assisted cell sorting strategies are established
to mitigate the non-specific uptake of immunomagnetic microparticles, enabling the
enrichment of endothelial cells to purities that prevent their overgrowth by stromal cells. This
work delineates a reliable method for acquiring human adipose tissue-derived microvascular
endothelial cells in large quantities with high purities that can be readily applied in future
vascular tissue engineering applications.
https://doi.org/10.1038/s42003-021-02732-8 OPEN
1Institute of Biomedical Engineering, University of Toronto, Toronto, ON, Canada. 2Translational Biology and Engineering Program, Ted Rogers Centre for
Heart Research, Toronto, ON, Canada. 3Department of Physiology, University of Toronto, Toronto, ON, Canada. 4Division of Plastic, Reconstructive, and
Aesthetic Surgery, University of Toronto, Toronto, ON, Canada. 5Departments of Surgery and Surgical Oncology, University Health Network, Toronto, ON,
Canada. 6Department of Mechanical and Industrial Engineering, University of Toronto, Toronto, ON, Canada. 7Faculty of Dentistry, University of Toronto,
Toronto, ON, Canada. ✉email: paul.santerre@utoronto.ca
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Comprising a single layer of cells lining the luminal surface
of the vasculature, the endothelium is the interface
between blood and the different tissues of the body1.
Although the endothelium exhibits marked phenotypic hetero-
geneity between vascular beds, its fundamental functions are to
maintain blood in a fluid state and to confine its flow to the
boundaries of the vasculature1. Endothelial cells (ECs) are con-
sequently indispensable for vascular tissue engineering, enabling
the vascularization and sustained perfusion of engineered tissues2,
as well as the endothelialization and prolonged patency of small-
diameter vascular prostheses3. Despite being the fundamental
building block for vascular tissue engineering, a clinically viable
source of endothelium has continued to elude the field4.
The ideal source of ECs for vascular tissue engineering will be
autologous to preclude immunogenic concerns and readily
accessible to minimize the time needed for their culture-mediated
expansion3. The scarcity of expendable blood vessels and the low
prevalence of microvascular ECs in tissues has prompted many to
turn to alternative sources of endothelium, namely stem and
progenitor cells5–7. However, their inherent proliferative and
differentiative potential, coupled with the spectrum of manip-
ulations used to instill in them an endothelial phenotype, intro-
duces distinct regulatory concerns that may impede their clinical
translation when compared with their natively differentiated and
minimally manipulated counterparts8,9. Remarkably, an abun-
dant and uniquely dispensable source of autologous endothelium
remains largely untapped: adipose tissue.
Adipose tissue is an attractive source of ECs for vascular tissue
engineering because it can be harvested autologously in large
quantities with minimally invasive procedures using a cannula
coupled to a liposuction pump or even a syringe10. While the
need for the culture-mediated expansion of human adipose
tissue-derived microvascular ECs (HAMVECs) is mitigated by
the abundant and uniquely dispensable nature of the tissue,
their low prevalence has continued to complicate their
acquisition3,11–15. Specifically, their primary cultures are often
overgrown by residual stromal cells from the cell sorting
procedure3,11–15. This challenge in acquiring HAMVECs may
account for their strikingly limited adoption for vascular tissue
engineering16–18. Moreover, the absence of an accessible alter-
native likely contributes to the sustained popularity of human
umbilical vein ECs (HUVECs), the use of which continues to be
reported in ~ 60% of publications amongst the biomaterials
and tissue engineering communities4,19 despite being an
allogeneic and, thereby, clinically impractical source of
endothelium20. Accordingly, there is a clear and unmet need for a
readily accessible and non-immunogenic source of endothelium if
tissue-engineered constructs are to leave the bench for the
bedside.
The overgrowth of microvascular ECs by residual stromal cells
from the cell sorting procedure is a common complication. This
phenomenon is not specific to HAMVECs3,11–15, but challenges
the isolation of microvascular ECs from nearly all tissues,
including the brain21, kidney22, and lung23. While early attempts
at their isolation relied on sieves and differential centrifugation to
enrich the endothelium from enzymatically digested tissues24,
these approaches have been largely replaced or augmented with
immunoselection, which allows for the enrichment of the endo-
thelium based on its unique cell-surface protein signature25.
Nevertheless, stromal cell contamination has continued to com-
plicate their primary cultures, driving many to resort to differ-
ential adhesion11,15,21, clonal selection23, and manual
weeding12,22 in order to prevent their impendent overgrowth.
These procedures are labour-intensive, time-consuming, and of
uncertain reproducibility, comprising a formidable obstacle to
their widespread adoption for vascular tissue engineering.
The objective of this study was to develop an accessible and
reliable method of acquiring endothelium from human fat for
vascular tissue engineering. Magnet-assisted cell sorting (MACS)
was explored for the immunoselection of HAMVECs due to its
low cost, small physical footprint, and ease of use. Their over-
growth by stromal cells was first investigated to gain a clear
understanding of the specific challenges underlying this phe-
nomenon, which was then used to inform the development of a
reproducible method of acquiring HAMVECs. Strategies were
established to mitigate the non-specific uptake of immunomag-
netic microparticles (IMPs), enabling the enrichment of HAM-
VECs to purities that prevent their overgrowth by stromal cells.
This study demonstrates the feasibility of sourcing autologous
endothelium from human fat for vascular tissue engineering. It
delineates a reliable and facile method for its acquisition that can
be readily implemented by the field.
Results
HAMVECs were often overgrown by residual ASCs from the
cell sorting procedure. HAMVECs were isolated from the stro-
mal vascular fraction of enzymatically digested human sub-
cutaneous abdominal white adipose tissue using MACS (Fig. 1).
The yield of stromal vascular cells was 6.6 ± 4.7 × 105cells per
gram of tissue. HAMVECs were extracted from the stromal
vascular fraction on the basis of a CD45−CD31+immunophe-
notype—i.e. a cell-surface protein signature characteristic of dif-
ferentiated endothelium26–28. The stromal vascular fraction was
first depleted of CD45+leucocytes prior to positively selecting for
CD31+HAMVECs due to the high prevalence of leucocytes
(41.8 ± 4.7% CD45+; Fig. 1a;Supplementary Fig. 1) and their
capacity to co-express this characteristic endothelial cell-surface
marker (43.0 ± 3.2% CD31+; Supplementary Fig. 2).
The putative CD45–CD31+HAMVECs comprised 0.9 ± 0.6%
of the stromal vascular fraction (Fig. 1a), which is comparable to
the prevalence of microvascular ECs in other tissues25. Their yield
was 4.6 ± 3.0 × 103cells per gram of fat. Cultures of HAMVECs
were significantly enriched for the CD45–CD31+immunophe-
notype when compared with the stromal vascular fraction
(98.6 ± 0.9% vs. 0.9 ± 0.6%, respectively; p< 0.0001; Fig. 1b), and
they exhibited a characteristic endothelial cobblestone-like
morphology (Fig. 1c). Cultures of these purities (98.6 ± 0.9%
CD45–CD31+; range: 98.0–99.7% CD45–CD31+) were success-
fully established from three patients to enable the ensuing
investigations of the challenges underlying their acquisition.
The isolation of HAMVECs had to be attempted from 20
patients in order to obtain these three cultures of a characteristic
endothelial cobblestone-like morphology. In the other 17 patients,
HAMVECs were visibly overgrown within 2 weeks by spindle-
shaped, fibroblast-like cells (Fig. 1g). These contaminating cells
comprised residual CD45−CD31−stromal vascular cells from the
MACS procedure (Fig. 1f). Importantly, sequential enrichments
for the CD31+immunophenotype failed to eliminate these
contaminating cells and prevent their overgrowth of HAMVECs
(Fig. 1g).
CD45−CD31−stromal vascular cells were retained from
subsequent isolations to facilitate investigations of their over-
growth of HAMVECs. They comprised 57.2 ± 5.0% of the stromal
vascular fraction (Fig. 1a), and their yield was 3.8 ± 2.7 × 105cells
per gram of tissue. Their cultures were significantly enriched for
the CD45−CD31−immunophenotype when compared with the
stromal vascular fraction (99.8 ± 0.2% vs. 57.2 ± 5.0%, respec-
tively; p=0.0019; Fig. 1d), and they exhibited a comparable
morphology to the contaminating cells (Fig. 1e). Notably, these
CD45−CD31−stromal vascular cells comprised adipose tissue-
derived stromal/stem cells (ASCs)29, having been previously
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validated to meet the phenotypic criteria delineated by the
International Federation for Adipose Therapeutics and Science and
the International Society for Cellular Therapy30.
HAMVECs were committed to the endothelial lineage. The
phenotypic heterogeneity of the endothelium and the paucity of
markers with specificity and sensitivity for the endothelial lineage
presents a challenge to the assessment of an endothelial
phenotype1. It has previously misled many to mistake monocytes
for endothelial progenitor cells31, platelets for circulating ECs32,
omental mesothelial cells for HAMVECs33,34, and even ASCs for
EC substitutes29. Accordingly, comprehensive phenotypic com-
parisons to representative EC controls were performed to confirm
the endothelial phenotype of the putative HAMVECs. Specifi-
cally, HAMVECs were compared to controls representative of the
predominant endothelial specializations35, namely arterial (con-
trol: human coronary artery ECs, HCAECs) vs. venous (control:
HUVECs) and macrovascular (controls: HCAECs and HUVECs)
vs. microvascular (control: human dermal microvascular ECs,
HDMVECs).
The endothelial phenotype of the putative HAMVECs was first
validated using a targeted assessment of characteristic endothelial
traits (Fig. 2). HAMVECs shared a similar cobblestone-like
morphology with the HUVECs, HCAECs, and HDMVECs
(Fig. 2a). The abundance of transcripts encoding CD31 (gene:
PECAM1), vascular endothelial (VE)-cadherin (gene: CDH5), and
von Willebrand Factor (vWF; gene: VWF) in HAMVECs was
statistically equivalent to that observed in the EC controls
(Fig. 2b; Supplementary Table 1; and Supplementary Fig. 3), and
they exhibited comparable expression of the corresponding
proteins (Fig. 2c). Furthermore, their uptake of acetylated low-
density lipoprotein (AcLDL) was similar (Fig. 2d). Importantly,
these morphological, molecular, and functional endothelial
hallmarks exhibited by HAMVECs were previously shown to be
negligible in ASCs when cultured under identical conditions and
compared with the same EC controls29. Lastly, HAMVECs
exhibited an angiogenic capacity comparable to that of the EC
Fig. 1 Primary cultures of human adipose tissue-derived microvascular endothelial cells (HAMVECs) are often overgrown by residual adipose tissue-
derived stromal/stem cells (ASCs) from the magnet-assisted cell sorting (MACS) procedure. The stromal vascular fraction of enzymatically digested
human subcutaneous abdominal white adipose tissue (a) was depleted of CD45+leucocytes prior to positively selecting for CD31 expression to establish
primary cultures of CD45−CD31+HAMVECs (b,c). Their primary cultures were often overgrown by residual CD45−
CD31−ASCs from the MACS
procedure despite sequential enrichments for CD31 expression (f,g), prompting the retention of ASCs for downstream studies (d,e). Shown are
representative pseudocolour plots depicting the composition of the different populations of cells (a,b,d,f), as well as representative photomicrographs
depicting their corresponding morphologies (c,e,g). Scale bars represent 200 µm; values mean ± standard deviation. While the isolation of HAMVECs was
initially attempted from twenty patients (n=20 biologically independent samples), only three cultures were visibly free of stromal cell overgrowth (n=3
biologically independent samples); in the other 17 patients, cultures were visibly overgrown by stromal cells within 2 weeks (n=17 biologically independent
samples). The composition of the different populations of cells was assessed in three patients in all but visibly contaminated primary cultures of HAMVECs
(a,b,d;n=3 biologically independent samples), in which flow cytometry was used to elucidate the identity of the contaminating stromal cells rather than
to assess their purity (f;n=1 biologically independent sample).
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controls (Fig. 2e), further supporting their functional endothelial
phenotype.
The commitment of HAMVECs to the endothelial lineage
was then assessed. Specifically, their adipogenic plasticity was
compared to that of ASCs (Fig. 3). While the culture of ASCs in
adipogenic media increased the size of their lipid droplets
(Fig. 3a), it did not significantly increase their total abundance of
lipids (Fig. 3b). This discrepancy may be attributed to differences
in their cellular densities, as the growth of ASCs was halted at the
confluence in adipogenic media but not in endothelial media
(Fig. 3a). Nevertheless, ASCs exhibited a significantly greater
accumulation of lipids than HAMVECs regardless of the media in
which they were cultured (Fig. 3b). Furthermore, the culture of
HAMVECs in adipogenic media induced their death rather than
adipogenesis (Fig. 3a), supporting that HAMVECs are fully
differentiated and committed to the endothelial lineage.
Proteomic assessment of HAMVECs implicated heterotypic
cell–cell interactions in the modulation of their overgrowth by
ASCs. The phenotype of HAMVECs was further compared to that
of the representative EC controls using liquid chromato-
graphy–tandem mass spectrometry (LC–MS/MS; Fig. 4). While
the endothelial proteome was 87% conserved (Fig. 4c), unsu-
pervised hierarchical clustering of their global proteomes
effectively grouped ≥2 biological replicates from each of the four
different vascular beds (Fig. 4b), supporting that the phenotypic
heterogeneity of endothelium persists despite its culture under
identical conditions35. HAMVECs exhibited the most distinctive
proteome (Fig. 4b, d), with 284 proteins differentiating them
from all other endothelial specializations. Importantly, gene
ontological analyses did not identify any biological pathways
related to angiogenesis, haemostasis, nor permeability, that were
enriched amongst these differentially expressed proteins, adding
further support to the endothelial phenotype of the CD45−
CD31+
stromal vascular cells. Rather, pathways implicated in proliferation
were found to be statistically overrepresented (Fig. 4e), suggesting
that the proliferative capacity of HAMVECs was their most dif-
ferentiating feature.
HAMVECs exhibited a significantly longer population dou-
bling time than HUVECs, HCAECs, and HDMVECs (Fig. 4f).
Interestingly, their slower proliferation was associated with the
slight but significantly lower purity of their cultures (Fig. 4g),
suggesting that residual ASCs from the cell sorting procedure
may suppress the proliferation of HAMVECs to facilitate their
overgrowth of primary cultures. However, HAMVECs could be
repeatedly sub-cultured and maintained at confluence for
>3 weeks without any signs of stromal cell overgrowth despite
their trace impurities. In fact, zones of inhibition appeared to
Fig. 2 Human adipose tissue-derived microvascular endothelial cells (HAMVECs) exhibit morphological, molecular, and functional hallmarks of
endothelium. HAMVECs were compared with endothelial cell (EC) controls representative of the predominant endothelial specializations, namely human
umbilical vein ECs (HUVECs; macrovascular, venous endothelium), human coronary artery ECs (HCAECs; macrovascular, arterial endothelium), and
human dermal microvascular ECs (HDMVECs; microvascular endothelium). aCobblestone-like morphology of endothelium. Scale bars represent 200 µm.
bAbundance of transcripts encoding CD31 (gene: PECAM1), vascular endothelial (VE)-cadherin (gene: CDH5), and von Willebrand Factor (vWF; gene:
VWF). Glyceraldehyde-3-phosphate dehydrogenase (gene: GAPDH) was used as a loading control. Dashed line depicts a mean difference of zero; and
dotted lines is the equivalence margin (∂) used for the two one-sided test for equivalence. Values represent the mean ± 90% confidence interval; mRNA,
messenger ribonucleic acid; and C
q
, the quantification cycle. cExpression and localization of the corresponding endothelial proteins. Scale bars represent
25 µm. dInternalization of acetylated low-density lipoprotein (AcLDL). Solid and dashed lines represent ECs cultured in the presence and absence of Alexa
Fluor 488-conjugated AcLDL, respectively. Values represent mean ± standard deviation. eCapillary-like tubulogenesis by ECs. Scale bars represent
200 µm. All experiments were performed in biological triplicate, using cells derived from three different donors (n=3 biologically independent samples).
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surround the residual stromal cells in these cultures (Fig. 4h),
suggesting that both HAMVECs and ASCs are capable of
suppressing each other’s proliferation.
HAMVECs and ASCs were seeded in pre-defined proportions
to recapitulate different efficacies of enrichment for the
CD45−CD31+immunophenotype, and the effect of seeding
purity on their proliferation and the consequent temporal
composition of their cultures was assessed (Fig. 5). HAMVECs
exhibited a significantly longer population doubling time than
ASCs (Fig. 5a), and their population growth rate was further
suppressed with lower seeding purities (Fig. 5b;Supplementary
Fig. 4). HAMVECs were also found to inhibit the growth of ASCs,
albeit to a significantly lower extent (Fig. 5b). Although similar
magnitudes of population growth inhibition in both HAMVECs
and ASCs were observed with a 90% seeding purity (Fig. 5b), the
longer population doubling time of HAMVECs (Fig. 5a) suggests
that an enrichment efficacy >90% is required in order for the
absolute population growth rate of HAMVECs to exceed that of
ASCs—i.e. the threshold of purity beyond which stromal cell
overgrowth is precluded. In fact, cultures of HAMVECs
established with 90% purity consistently exhibited significant
overgrowth by ASCs within 7 days, and HAMVECs were virtually
undetectable in these cultures after 28 days (Fig. 5c). In contrast,
our isolates (98.6 ± 0.9% CD45−CD31+; range: 98.0–99.7%
CD45−CD31+) could be maintained in culture for 28 days
without a significant decline in purity (Fig. 5c). These findings
underscore the importance of seeding purity to the protracted
stability of HAMVEC cultures, and suggest that no further
enrichment is needed in cultures established with purities ≥98%.
ASCs exhibited a capacity to bind and internalize anti-CD31
IMPs. The failure of sequential rounds of MACS to eliminate
contaminating ASCs from primary cultures suggests that they are
capable of binding the anti-CD31 IMPs. Interestingly, ASCs
exhibited both a membrane-bound and intracellular localization
of the anti-CD31 IMPs (Fig. 6a), indicating that they can further
internalise them. The prevalence of ASCs exhibiting a membrane-
bound or intracellular localization of anti-CD31 IMPs after
20 min in suspension (i.e. labelling conditions for MACS) was
17.1 ± 3.3% (Fig. 6b;Supplementary Fig. 5), supporting a role for
the IMPs in mediating the contamination of primary cultures.
While the prevalence of membrane-bound anti-CD31 IMPs
decreased over time in culture, the prevalence of their inter-
nalization significantly increased (Fig. 6b; p< 0.0001). In fact, the
proportion of ASCs exhibiting bound and internalized anti-CD31
IMPs was significantly greater after 48 h in culture than after
20 mins in suspension (Fig. 6c), accounting for the failure of
sequential rounds of MACS to eliminate the ASCs. Moreover,
IMP-laden ASCs exhibited comparable levels of deoxyribonucleic
acid (DNA) synthesis as IMP-free ASCs (Fig. 6d;Supplementary
Fig. 6), suggesting that their binding and internalization of the
anti-CD31 IMPs does not significantly affect their capacity to
proliferate and overtake HAMVECs following their magnetic
separation and sub-culture.
Uptake of anti-CD31 IMPs by ASCs was non-specific, but size
and exposure -dependent. The capacity of ASCs to express low
levels of characteristic endothelial markers is well-documented29,
suggesting that their binding and internalization of the anti-CD31
IMPs may be a manifestation of their limited endothelial plasti-
city. LC–MS/MS was used to identify an alternative immuno-
phenotypic marker with greater specificity for HAMVECs in
cultures contaminated with ASCs (Fig. 7). While unsupervised
hierarchical clustering of their global proteomes supported the
distinct phenotypes of HAMVECs and ASCs (Fig. 7b), 88% of
their proteomes were conserved (Fig. 7c). HAMVECs were found
to express all of the characteristic immunophenotypic markers
traditionally used to define ASCs30, including CD90, CD44,
CD13, CD73, CD29, and CD105 (Fig. 7d). Furthermore, the
endothelial plasticity of the ASCs was evident, with LC–MS/MS
detecting their expression of CD31, VE-cadherin, and vWF, albeit
in significantly lower abundances when compared with the
HAMVECs (Fig. 7d). These findings suggest that HAMVECs
cannot be enriched, nor can ASCs be depleted, from primary
cultures on the basis of their conventional immunophenotypes—
including CD31. Gene ontological mapping of the 457 proteins
enriched in HAMVECs to their cellular components identified
188 that were localized to the plasma membrane (Supplementary
Fig. 7), of which 37 were detected with specificity and sensitivity
(i.e. n=3/3 HAMVECs and n=0/3 ASCs). CD93 was selected as
Fig. 3 Adipogenic plasticity is evident in adipose tissue-derived stromal/stem cells (ASCs), not human adipose tissue-derived microvascular
endothelial cells (HAMVECs). HAMVECs and ASCs were cultured in adipogenic medium for 10 days before their accumulation of lipids was assessed by
Oil Red O with hematoxylin counterstaining. Endothelial medium is used as control. Shown are representative photomicrographs depicting the
accumulation of lipids by HAMVECs and ASCs (a), as well as a bar graph delineating its quantification (b). Hematoxylin stains nuclei blue/purple, and Oil
Red O stains lipids red/orange. Scale bars represent 100 µm. Values represent mean ± standard deviation; and *p< 0.05. Experiments were performed in
biological triplicate, using cells derived from three different donors (n=3 biologically independent samples).
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a candidate immunophenotypic marker for HAMVECs due to its
cell-surface localization and prior characterization36.
The specificity and sensitivity of CD93 for HAMVECs was
validated both in culture as well as in the freshly isolated stromal
vascular fraction. While the sensitivity of CD93 for HAMVECs
in vitro was slightly poorer than that of CD31 (HAMVECs:
99.6 ± 0.4% CD31+vs. 95.2 ± 4.3% CD93+;p=0.2244), its
specificity was greater (ASCs: 8.4 ± 5.8% CD31+vs. 0.1 ± 0.0%
CD93+;p=0.1280; Fig. 7e). Although these differences were not
statistically significant and the detection of CD31 in ASCs
required its secondary antibody-mediated signal amplification
(Supplementary Fig. 8), these slight disparities in sensitivity and
specificity may have meaningful implications on their capacities
to enrich for HAMVECs given the potent growth inhibition
imposed by even a small number of residual ASCs. Interestingly,
the specificity and sensitivity exhibited by CD93 for HAMVECs
in vitro were not shared by HAMVECs in the stromal vascular
fraction, with low levels of CD93 expression detected amongst
CD45+leucocytes, CD45−CD31−ASCs, and CD45–CD31+
HAMVECs (Fig. 7f). These findings suggest that although
CD93 cannot be used to isolate HAMVECs from the stromal
vascular fraction of enzymatically digested adipose tissue, it may
be superior to CD31 for the sequential enrichment of their
contaminated cultures.
The binding and internalization of anti-CD93 IMPs by ASCs
was compared to that of the anti-CD31 IMPs. Anti-CD93
Fig. 4 Proteomic assessment of human adipose tissue-derived microvascular endothelial cells (HAMVECs) suggests that their proliferation is
suppressed by adipose tissue-derived stromal/stem cells (ASCs). a Workflow depicting the proteomic comparison of HAMVECs with endothelial cell
(EC) controls representative of the predominant endothelial specializations, namely human umbilical vein ECs (HUVECs; macrovascular, venous
endothelium), human coronary artery ECs (HCAECs; macrovascular, arterial endothelium), and human dermal microvascular ECs (HDMVECs;
microvascular endothelium). bUnsupervised hierarchical clustering of the proteomes of the ECs derived from four different vascular beds. cDistribution of
detected proteins amongst the different ECs. dHierarchical clustering of the proteins detected in different abundances (p< 0.05) between EC types.
eBiological pathways differentiating HAMVECs from all other types of ECs. Black bars highlight those related to proliferation; and white bars metabolism.
fPopulation doubling times of the different ECs, determined from their exponential growth phase observed over 7 days of culture. Values represent
mean ± standard deviation; and *p< 0.05 compared with HAMVECs. gPurities of the different EC cultures. Values represent mean ± standard deviation;
and, *p< 0.05 compared with HAMVECs. hRepresentative zones of inhibition surrounding residual ASCs in primary cultures of HAMVECs that were
maintained at confluence for over 3 weeks. Scale bar represents 500 µm. All experiments were performed in biological triplicate, using cells derived from
three different donors (n=3 biologically independent samples).
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antibodies were conjugated to superparamagnetic microparticles
through a DNA linker, generating cleavable (c)IMPs. These anti-
CD93 cIMPs were slightly but significantly larger in modal
diameter than the commercial anti-CD31 IMPs (4.8 ± 0.5 µm vs.
4.4 ± 0.6 µm, respectively; p< 0.0001), prompting the generation
of anti-CD31 cIMPs of a comparable size (4.8 ± 0.6 µm;
p=0.3772 compared with anti-CD93 cIMPs; Fig. 7g). The
binding and internalization of anti-CD93 cIMPs by ASCs were
not significantly different from that of the anti-CD31 cIMPs after
20 min in suspension nor 48 hr in culture, although the
internalization of both (anti-CD31 and anti-CD93) cIMPs was
significantly lower than that of the smaller anti-CD31 IMPs after
48 hr in culture (Fig. 7h). These findings suggest that the binding
and internalization of IMPs by ASCs is non-specific, but is
dependent on the size of the microparticles and their exposure to
the cells.
The effect of IMP size on their uptake by ASCs was further
explored (Fig. 8;Supplementary Fig. 9). Five distinct anti-CD31
IMPs ranging in modal diameter from 0.9 ± 0.3 µm to
8.7 ± 1.5 µm were investigated (Fig. 8a;Supplementary Table 2).
There was an inverse relationship between IMP size and their
uptake by ASCs (Fig. 8b; p< 0.0001). Interestingly, IMP size also
had a significant effect on the temporal dynamics underlying their
uptake (Fig. 8b; p=0.0003). While the uptake of anti-CD31
Fig. 5 Heterotypic cell–cell interactions modulate the overgrowth of human adipose tissue-derived microvascular endothelial cells (HAMVECs) by
adipose tissue-derived stromal/stem cells (ASCs). a Population doubling times of HAMVECs and ASCs, determined from their exponential growth phase
observed over 7 days of culture. *p< 0.05 compared with HAMVECs. bEffect of seeding purity on the population growth rates of HAMVECs and ASCs,
observed over 4 days of culture. *p< 0.05 compared with HAMVECs; #p< 0.05 compared with 100%. cEffect of seeding purity on the composition of
cultures maintained at confluence for up to 28 days. *p< 0.05 compared with 100% seeding purity at respective time-point; #p< 0.05 compared with
purity at day 1. Values represent mean ± standard deviation. All experiments were performed in biological triplicate, using cells derived from three different
donors (n=3 biologically independent samples).
Fig. 6 Adipose tissue-derived stromal/stem cells (ASCs) bind and internalize anti-CD31 immunomagnetic microparticles (IMPs). a Intracellular
localization of anti-CD31 IMPs in ASCs after 24 hr of culture. Scale bar represents 25 µm. bPrevalence of IMP localization in ASCs after 20 min in
suspension (i.e. labelling conditions for magnet-assisted cell sorting procedure) and different durations in culture. *p< 0.05 compared with respective
localization in suspension; #p< 0.05 compared with membrane-bound localization. cTotal prevalence of ASC interactions with anti-CD31 IMPs after
20 min in suspension vs. 48 hr in culture. Shown is a stacked bar graph depicting the total prevalence of ASCs exhibiting a membrane-bound and/or
intracellular localization of anti-CD31 IMPs, and the corresponding proportions of each. Values represent mean ±standard deviation of the total prevalence.
Triangles depict an internalized localization, and diamonds depict a membrane-bound localization. *p< 0.05 compared with suspension. dIncorporation of
thymidine analogue 5-ethynyl-2'-deoxyuridine (EdU) during deoxyribonucleic acid (DNA) synthesis by IMP-laden and IMP-free ASCs after 48 hr in culture.
ns represents not statistically significant. All values represent mean ± standard deviation. All experiments were performed in biological triplicate, using cells
derived from three different donors (n=3 biologically independent samples).
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IMPs ≥4.4 µm in diameter was exacerbated in culture, those that
were ≤3.9 µm in diameter were readily taken up by ASCs after
20 min in suspension, and their culture did not significantly
increase their uptake (Fig. 8b). This may be attributed to a greater
capacity for ASCs to bind and retain smaller IMPs at the plasma
membrane prior to their internalization (Supplementary Fig. 9e).
These findings suggest that IMPs ≤3.9 µm in diameter cannot be
used to isolate HAMVECs due to the capacity for ASCs to readily
bind them during the labelling portion of the MACS procedure
(i.e. 20 min in suspension), and that the non-specific uptake of
anti-CD31 IMPs ≥4.4 µm in diameter can be mitigated by either
limiting their exposure to the cells to 20 min in suspension or by
using superparamagnetic microparticles of a larger diameter.
Mitigating the non-specific uptake of IMPs enabled the
enrichment of HAMVECs. The acquisition of HAMVECs by
MACS is a two-step procedure: the first step involves the isola-
tion of HAMVECs from the stromal vascular fraction of enzy-
matically digested fat in order to establish their primary cultures,
and the second step involves the enrichment of the primary
Fig. 7 Uptake of immunomagnetic microparticles (IMPs) by adipose tissue-derived stromal/stem cells (ASCs) is non-specific, but size and exposure
dependent. a Workflow depicting the proteomic comparison of human adipose tissue-derived microvascular endothelial cells (HAMVECs) and ASCs used
to identify immunophenotypic markers with specificity for HAMVECs, which were then validated both in vitro and in situ before generating IMPs and
evaluating their binding and internalization by ASCs. bUnsupervised hierarchical clustering of the global proteomes of HAMVECs and ASCs. cDistribution
of proteins detected in HAMVECs and ASCs. dRelative abundances of proteins detected in both HAMVECs and ASCs. eCell-surface expression of CD31
and CD93 by HAMVECs and ASCs in culture. Solid lines represent stained cells; dashed lines, isotype controls. fSpecificity and sensitivity of CD93 for
HAMVECs in the stromal vascular fraction of enzymatically digested human subcutaneous abdominal white adipose tissue. Presented is a t-distributed
stochastic neighbour embedding (tSNE) plot depicting the unsupervised clustering of stromal vascular cells based on their cell-surface expression of CD45,
CD31, and CD93, as well as histograms depicting the cell-surface expression of CD93 amongst the three principal subpopulations: CD45−CD31+
HAMVECs, CD45−CD31−ASCs, and CD45+leucocytes. Solid lines represent stained cells; and dashed lines, fluorescence minus one controls. gSize
distributions of IMPs. Commercial anti-CD31 IMPs were compared with cleavable (c)IMPs generated to be reactive against CD31 or CD93. *p< 0.05
compared with anti-CD31 IMPs. hBinding and internalization of IMPs by ASCs after 20 min in suspension (i.e. labelling conditions for magnet-assisted cell
sorting procedure) and 48 hr in culture. Shown is a stacked bar graph depicting the total prevalence of ASCs exhibiting a membrane-bound and/or
intracellular localization of the IMPs, and the corresponding proportions of each. Values represent mean ± standard deviation of the total prevalence.
Triangles depict an internalized localization, and diamonds depict a membrane-bound localization.*p< 0.05 compared with anti-CD31 IMPs. e,hValues
represent mean ± standard deviation. All experiments were performed in biological triplicate, using cells derived from three different donors (n=3
biologically independent samples).
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cultures in order to eliminate any residual, contaminating ASCs.
The limited endothelial plasticity of ASCs and their capacity to
bind and internalize IMPs were hypothesized to undermine the
enrichment of contaminated cultures of HAMVECs. Our pre-
ceding experiments suggest that the uptake of anti-CD31 IMPs by
ASCs is non-specific, but size and exposure-dependent. Hence,
the effects of alternative target antigens (CD31 vs. CD93), sizes
(4.4 µm vs. 4.8 µm), and exposures (suspension vs. culture) of
IMPs on the enrichment efficacy of the MACS procedure were
investigated using an in vitro model of contaminated primary
cultures (Fig. 9a). Specifically, HAMVECs and ASCs were seeded
in a 9:1 proportion with or without IMPs to recapitulate their
potential exclusion from cultures, made possible by enzymatically
cleaving the DNA linkers coupling the antibodies to the super-
paramagnetic microparticles. Their enrichment was performed
after 4 days, and their purities (% CD31+) were evaluated after
7 days (Fig. 9b;Supplementary Fig. 10). In the absence of their
enrichment, the purity of the co-cultures deteriorated to
40.0 ± 26.0% (Fig. 9c), underscoring the need for seeding purities
>90% for the absolute population growth rate of HAMVECs to
exceed that of ASCs.
MACS was effective in increasing the purity of the co-cultures,
and there was no significant difference between enriching for
HAMVECs on the basis of their expression of CD31 and CD93
(Fig. 9c). Moreover, there was no significant difference between
the enrichment of IMP-free cultures and those laden with the
larger 4.8 µm (anti-CD31 and anti-CD93) cIMPs, but their
enrichment was more effective than that of cultures laden with
the smaller 4.4 µm (anti-CD31) IMPs (Fig. 9d). Although not
statistically significant, the enrichment of IMP-free cultures
yielded higher purities with less variability than the enrichment
of 4.8 µm (anti-CD31 and anti-CD93) cIMP-laden cultures
(98.9 ± 0.7% vs. 91.0 ± 8.2%, respectively; p=0.4612; Fig. 9d),
reflecting the greater capacity of ASCs to internalize 4.8 µm (anti-
CD31 and anti-CD93) cIMPs in culture than in suspension
(8.1 ± 5.0% after 48 hr in culture vs. 0.2 ± 0.2% after 20 min in
suspension; p=0.0123; Fig. 7h). These findings support the non-
specific but size and exposure-dependent uptake of IMPs by
ASCs, and indicate that the limited capacity of ASCs to express
CD31 does not necessitate the utilization of an alternative cell-
surface marker with greater specificity for cultured HAMVECs.
By mitigating the introduction of ASCs into primary cultures and
by enabling their effective sequential enrichment, the DNase-
mediated exclusion of anti-CD31 cIMPs from cultures has been
used to reliably acquire HAMVECs with high purities from an
additional five consecutive patients (98.7 ± 0.5% CD45−CD31+;
n=5).
Discussion
Adipose tissue is an attractive source of ECs for vascular
tissue engineering because it can be harvested autologously in
large quantities with minimally invasive procedures10. While
the need for the culture-mediated expansion of HAMVECs is
mitigated by the abundant and uniquely dispensable nature of
the tissue, the low prevalence of HAMVECs has continued to
complicate their acquisition, with their primary cultures often
being readily overgrown by fibroblast-like stromal cells3,11–15.
Here, we demonstrate that the non-specific uptake of IMPs
by these residual ASCs from the cell sorting procedure under-
mines the efficacy of sequential enrichments for HAMVECs.
The non-specific uptake of IMPs can be mitigated through
the use of superparamagnetic microparticles of a larger diameter
or by excluding them from primary cultures where they can
be more readily internalized by ASCs. Both of these strategies
can mitigate the non-specific uptake of IMPs and can be easi-
ly implemented to facilitate the reliable acquisition of HAMVECs
in large quantities with high purities for vascular tissue
engineering.
MACS was used for the immunoselection of HAMVECs due to
its accessibility. Its low cost, small physical footprint, and ease of
use make it more amenable for a number of stakeholders when
compared with alternatives such as fluorescence-assisted cell
sorting. This is an important consideration in promoting the
clinical translation of tissue-engineered products and is under-
scored by the sustained popularity of HUVECs amongst the
biomaterials and tissue engineering communities4,19 in the
absence of a readily accessible and clinically viable alternative.
The use of IMPs for both the isolation and sequential enrichment
of HAMVECs is not only accessible, but may also be more
scalable and less variable than other techniques such as differ-
ential adhesion11,15,21, clonal selection23, and manual
weeding12,22, which are labour-intensive and susceptible to
human error in discriminating endothelium from stromal cells.
Immunoselection eliminates human error in the discrimination
of cell types by exploiting their cell-surface protein signatures.
The enrichment of HAMVECs was pursued rather than the
Fig. 8 Size affects the extent and temporal dynamics underlying the
uptake of immunomagnetic microparticles (IMPs) by adipose tissue-
derived stromal/stem cells (ASCs). a Size distribution of five distinct anti-
CD31 IMPs. *p< 0.05 compared with all other IMPs. bTotal uptake (i.e.
membrane-bound and/or internalized) of the different anti-CD31 IMPs by
ASCs after 20 min in suspension (i.e. labelling conditions for magnet-
assisted cell sorting) and 48 hr in culture. Values represent
mean ± standard deviation; *p< 0.05; and, ns represents not significant.
This experiment was performed in biological triplicate, using cells derived
from three different donors (n=3 biologically independent samples).
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depletion of non-endothelial cell types due to the heterogeneity of
the stromal vascular fraction. Immunophenotyping revealed that
HAMVECs account for only 1% of the stromal vascular fraction,
with the remainder comprising leucocytes and ASCs. Single-cell
RNA sequencing has highlighted that the heterogeneity of the
stromal vascular fraction is even greater, with each of these three
groups of cells being comprised of multiple subpopulations37. The
challenge in identifying a cell-surface protein signature that
encompasses the phenotypic heterogeneity of non-endothelial cell
types may account for the failure of negatively selecting against
CD14 and F11 to obtain HAMVECs with purities >90%13,14.
The purity of HAMVECs is of marked importance to the
protracted stability of their cultures. While the faster rate of
proliferation of ASCs and their potent growth inhibition of
HAMVECs facilitates their overgrowth of primary cultures,
HAMVECs induce a reciprocal, albeit weaker, suppression of
ASC proliferation. Therefore, the overgrowth of primary cultures
may be prevented by achieving a threshold of purity beyond
which the absolute population growth rate of HAMVECs exceeds
that of ASCs. Although Arts et al. have suggested that a purity
≥75% is sufficient, this conclusion was based on the thicknesses
of tissues generated after 3 weeks of co-culturing HUVECs and
human foetal lung fibroblasts in several pre-defined
proportions14. In contrast, our co-cultures of HAMVECs and
ASCs exhibited a deterioration in purity from 90 to 40% after
only 1 week. This discrepancy may be attributed to the differential
rates of proliferation and disparate cell–cell interactions between
HUVECs and human foetal lung fibroblasts vs. HAMVECs and
ASCs. Our primary cultures of HAMVECs could be repeatedly
sub-cultured and maintained at confluence for 4 weeks without
Fig. 9 Mitigating the non-specific uptake of immunomagnetic microparticles (IMPs) by adipose tissue-derived stromal/stem cells (ASCs) facilitates
the enrichment of human adipose tissue-derived microvascular endothelial cells (HAMVECs). a Schematic depicting the magnet-assisted cell sorting
(MACS) procedure used to acquire HAMVECs from enzymatically digested fat, as well as the different combinations of IMPs tested for the isolation and
enrichment of HAMVECs using an in vitro model of their contaminated primary cultures. Specifically, HAMVECs and ASCs were seeded in a 9:1 proportion
with or without IMPs to recapitulate their potential exclusion from primary cultures, made possible by enzymatically cleaving the deoxyribonucleic acid
linkers coupling the antibodies to the superparamagnetic microparticles (i.e. cleavable (c)IMPs). Their enrichment was performed after 4 days, and their
purities (% CD31+) were evaluated after a total of 7 days. Created with BioRender.com bEffects of alternative target antigens (CD31 vs. CD93), sizes
(4.4 µm IMPs vs. 4.8 µm cIMPs), and exposures (introduced vs. excluded from cultures) of IMPs on the enrichment efficacy of the MACS procedure.
Values represent mean. cEffect of the target antigen on the enrichment efficacy of the MACS procedure. ‘None’depicts the distribution of purities of co-
cultures that were not subjected to MACS after 4 days; ‘CD31’, those that were enriched using anti-CD31 IMPs or anti-CD31 cIMPs; and, ‘CD93’, those that
were enriched using anti-CD93 cIMPs. dEffect of introducing IMPs into primary cultures on the efficacy of their sequential enrichment. ‘None’depicts the
distribution of purities of enriched co-cultures that were free of IMPs and cIMPs at the time of MACS; ‘4.4 µm IMPs’, those that were laden with the
smaller anti-CD31 IMPs; and, ‘4.8 µm cIMPs’, those that were laden with the larger anti-CD31 cIMPs or anti-CD93 cIMPs. *p< 0.05; ns represents not
statistically significant. Dashed lines in the violin plots represent median; dotted lines, quartiles; and, horizontal solid lines, range. The experiment was
performed in biological triplicate, using cells derived from three different donors (n=3 biologically independent samples).
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any signs of stromal cell overgrowth if established with
purities ≥98%.
The paucity of markers with specificity and sensitivity for the
endothelial lineage complicated the acquisition of HAMVECs.
The stromal vascular fraction had to be depleted of CD45+leu-
cocytes prior to positively selecting for CD31+HAMVECs due to
their capacity to co-express this characteristic endothelial marker.
Furthermore, contaminating ASCs from the cell sorting proce-
dure were found to express characteristic endothelial markers,
including CD31, VE-cadherin, and vWF. Although the temporal
dynamics underlying the expression of these conventional phe-
notypic markers were not investigated, the differential detection
of CD93 in vitro and in situ supports that the action of culture
itself induces a phenotypic change in both ECs38 and ASCs39.
However, the extent to which this phenomenon is a manifestation
of their phenotypic plasticity versus the culture-mediated selec-
tion for their different subpopulations remains unclear.
The phenotypic heterogeneity of the stromal vascular fraction
is not limited to ASCs and leucocytes, but is also evident amongst
the ECs37. CD31 is a pan-endothelial marker1, suggesting that
HAMVECs isolated from the stromal vascular fraction on the
basis of a CD45−CD31+immunophenotype comprise a hetero-
geneous mixture of arterial, venous, and lymphatic subpopula-
tions. Interestingly, CD93 has been reported to preferentially
stain post-capillary venules36, suggesting that its greater sensi-
tivity for HAMVECs in vitro than in situ may reflect the culture-
mediated selection for the venous subpopulation. The phenotypic
heterogeneity of CD45−CD31+HAMVECs is an important area
for further research as it may mitigate the phenotypic hetero-
geneity observed between ECs derived from different vascular
beds1,35; i.e., the putatively heterogeneous CD45−CD31+HAM-
VECs may be further enriched for desirable phenotypic specia-
lizations for tailored vascular tissue engineering applications3.
The capacity of anti-CD31 IMPs to establish primary cultures
of HAMVECs but not facilitate their subsequent enrichment was
believed to have been the manifestation of the de novo expression
of CD31 by ASCs induced by their culture in medium containing
vascular endothelial growth factor and basic fibroblast growth
factor29. However, despite detecting CD31 in cultured ASCs by
LC–MS/MS and validating its cell-surface localization by flow
cytometry, its efficacy as a target antigen for the enrichment of
HAMVECs was comparable to that of CD93. The comparable
enrichment efficacies of anti-CD31 IMPs and anti-CD93 IMPs
despite the greater specificity of CD93 for cultured HAMVECs
may be attributed to the low level of expression of CD31 by ASCs.
The abundance of CD31 in ASCs was <0.08% of that in HAM-
VECs, requiring secondary antibody-mediated signal amplifica-
tion of the directly conjugated anti-CD31 antibody to facilitate its
detection by flow cytometry. The binding affinity of antibodies is
dependent on the antigen density present on the cell surface40,
suggesting that the cell-surface density of CD31 amongst ASCs
may have been too sparse to facilitate their stable binding of the
anti-CD31 IMPs.
The sequential enrichment of primary cultures was under-
mined by the non-specific uptake of the anti-CD31 IMPs. The
internalization of particles is dependent on not only their size,
shape, and surface chemistry41, but also the cell type42. While the
capacity for leucocytes to uptake micron-sized particles is well-
established43, their internalization by other cell types has been
largely dismissed44. The commercial anti-CD31 IMPs utilized in
this investigation were 4.4 µm in diameter (volume: 45 µm3) and
were readily internalized by over 25% of ASCs. Subsequent
enrichments for CD31 expression consequently selected for not
only HAMVECs, but also the ASCs that had internalized the anti-
CD31 IMPs. While the non-specific internalization of the anti-
CD31 IMPs could be mitigated through the use of microparticles
of a larger diameter (e.g. diameter: 4.8 µm; volume: 58 µm3), their
exclusion from cultures, made possible by enzymatically cleaving
DNA linkers coupling the antibodies to the superparamagnetic
microparticles, was found to be most effective in facilitating the
acquisition of HAMVECs with the highest purity and least
variability.
The low prevalence of microvascular ECs in tissues has
remained a formidable obstacle to their reliable acquisition,
prompting many to turn to alternative sources of endothelium for
vascular tissue engineering at the expense of increased regulatory
scrutiny. The challenges underlying their isolation and expansion
were investigated to develop an accessible and reliable method of
obtaining them from human fat—an abundant and uniquely
dispensable source of autologous endothelium for the vascular-
ization of tissue-engineered constructs and the endothelialization
of small-diameter vascular prostheses. Although disparate growth
kinetics and the paucity of markers with specificity and sensitivity
for the endothelial lineage challenged their acquisition, mitigating
the non-specific uptake of IMPs was imperative for the effective
sequential enrichment of HAMVECs to purities that prevented
their overgrowth by ASCs. The findings of this study demonstrate
the feasibility of sourcing autologous endothelium from human
fat, and delineate a reliable and facile method for its acquisition
from patients that can be readily applied in future vascular tissue
engineering applications.
Methods
Materials. Subcutaneous abdominal white adipose tissue was obtained with
informed consent from 25 patients undergoing reconstructive breast surgery at the
University Health Network (Toronto, ON, Canada; institutional research ethics
board approval no. 13-6437-CE). Tissue culture-treated polystyrene (TCPS) was
sourced from Corning (Corning, NY, United States). Unless indicated otherwise, all
other materials were from Sigma-Aldrich (St. Louis, MO, United States).
Isolation of the stromal vascular fraction. The stromal vascular fraction was
isolated from adipose tissue as previously described29. Briefly, adipose tissue was
minced and enzymatically digested for 1 hr at 37 °C using collagenase type II
(2 mg/mL) in Kreb’s Ringer bicarbonate buffer supplemented with 3 mM glucose,
25 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid, and 20 mg/mL bovine
serum albumin (BSA). The buoyant adipocytes were discarded following the
centrifugation of the digest, and the pelleted tissue was subjected to another 15 min
of enzymatic digestion at 37 °C using 0.25% trypsin-ethylenediaminetetraacetic
acid (EDTA) solution. The cells were then resuspended in sterile distilled and
deionized water supplemented with 0.154 M ammonium chloride, 10 mM potas-
sium bicarbonate, and 0.1 mM EDTA for 10 min to facilitate erythrocyte lysis,
before being filtered through a 100 µm sieve. The resulting filtrate was defined as
the stromal vascular fraction and was immediately prepared for MACS or
immunophenotyping.
Magnet-assisted cell sorting. IMPs directed against CD45 (Dynabeads 45) and
CD31 (Dynabeads CD31 Endothelial Cell) were obtained from Invitrogen
(Carlsbad, CA, United States). Alternatively, cIMPs were generated using the
CELLection Biotin Binder Kit (Invitrogen). Specifically, 5 µg of either mouse anti-
human CD31 antibodies (Miltenyi Biotec, Bergisch Gladbach, Germany; catalogue
no. 130-119-893) or mouse anti-human CD93 antibodies (BioLegend, San
Diego, CA, United States; catalogue no. 336104) were conjugated to 107Dynabeads
via biotin-streptavidin interactions through DNA linkers, enabling their release
from cells with the supplied DNase.
HAMVECs and ASCs were isolated from the stromal vascular fraction using
MACS. Cells were resuspended in sorting buffer (phosphate-buffered saline
without calcium chloride and magnesium chloride (PBS−/−) supplemented with
2 mM EDTA and 0.1% (w/v) BSA), and were incubated with IMPs for 20 min at
4 °C before being magnetically separated using the DynaMag-5 Magnet
(Invitrogen). The stromal vascular fraction was depleted of CD45+leucocytes prior
to separating CD45−CD31+HAMVECs from CD45−CD31−ASCs. The
enzymatic exclusion of cIMPs from cultures was performed in select experiments
using the supplied DNase as per the manufacturer’s instructions. Briefly, cIMP-
labelled cells were incubated with DNase to dissociate the cells from the
superparamagnetic microparticles prior to using the DynaMag-5 Magnet to
separate the superparamagnetic microparticles from the cells of interest. The latter
were then seeded onto TCPS in the absence of the superparamagnetic
microparticles used to isolate them. Cultures of HAMVECs were enriched using
IMPs or cIMPs directed against CD31 or CD93 using the same procedure.
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Cell culture. HAMVECs and ASCs were plated onto TCPS and cultured at 37 °C,
5% CO
2
under a relative humidity of 85% in Endothelial Cell Growth Medium
(EGM)-2 (Lonza, Walkersville, MD, United States). HUVECs (Lonza; n=3 bio-
logically independent samples), HCAECs (Lonza; n=3 biologically independent
samples), and HDMVECs (Lonza and PromoCell, Heidelberg, Baden-Württem-
berg, Germany; n=3 biologically independent samples) were obtained commer-
cially and cultured under the same conditions. Media was exchanged three times a
week, and cells were passaged at 75–90% confluence using TrypLE Express
(Invitrogen). Phase-contrast transmission light microscopy was used to assess
morphology and confluence (Leica DMIL, Leica Microsystems, Wetzlar, Hesse,
Germany). Cells were counted using a hemocytometer, and dead cells were
excluded based on trypan blue staining.
Immunophenotyping. Cells were stained with either Live/Dead Fixable Aqua or
Live/Dead Fixable Near-IR (Invitrogen), and Fc receptors were blocked using
Human TruStain FcX (BioLegend). Cells were then stained for 20 min at 4 °C with
combinations of the following fluorophore-conjugated mouse anti-human mono-
clonal antibodies sourced from BioLegend: CD45-APC/Cy7 (catalogue no.
368516), CD45-Brilliant Violet 785 (catalogue no. 304048), CD31-Alexa Fluor 488
(catalogue no. 303110), CD31-Brilliant Violet 421 (catalogue no. 303124), and
CD93-PE (catalogue no. 336108). The staining concentration for each of the
antibodies was 5 µg/mL. For select experiments, the signal of the Alexa Fluor 488-
conjugated mouse anti-human CD31 antibody (BioLegend; catalogue no. 303110;
staining concentration, 5 µg/mL) was amplified using an Alexa Fluor 488-
conjugated goat anti-mouse IgG secondary antibody (Invitrogen; catalogue no.
A11001; staining concentration, 5 µg/mL; Supplementary Fig. 8). Stained cells were
fixed with 4% (w/v) paraformaldehyde in PBS−/−for 15 min at 4 °C. Compensa-
tion was achieved using the AbC Anti-Mouse Bead Kit and the ArC Amine
Reactive Compensation Bead Kit (Invitrogen). Gates were set using fluorescence
minus one controls. Flow cytometry was performed using a BD LSR II or BD LSR
Fortessa flow cytometer (Becton, Dickinson and Company, Franklin Lakes, NJ,
United States) at the Temerty Faculty of Medicine Flow Cytometry Facility
(University of Toronto, Toronto, ON, Canada). Data was acquired using BD
FACSDiva software version 8.0.1, and analysed using FlowJo software version
10.7.1 (Becton, Dickinson and Company). The immunophenotypes of two popu-
lations of cells were compared using two-tailed t-tests, blocking for donors when
appropriate; and, those of ≥3 populations were compared using a one-way analysis
of variance (ANOVA), with Tukey’s post-hoc test for multiple comparisons.
Gene expression. Reverse transcription quantitative real-time polymerase chain
reaction was performed as previously described29. Briefly, total ribonucleic acid
isolated from cells using Trizol (Invitrogen) was immediately reverse transcribed
into complementary DNA (cDNA) using the High Capacity cDNA Reverse
Transcription Kit (Applied Biosystems, Foster City, CA, United States). The
quantitative real-time polymerase chain reaction was performed on a CFX384
Touch Real-Time PCR Detection System (Bio-Rad, Hercules, CA, United States),
using the SsoAdvanced Universal SYBR Green Supermix (Bio-Rad). Each reaction
comprised 10 ng template cDNA and 450 nM of both forward and reverse primers
in a total volume of 10 µL, with thermal cycling performed as previously
described29. The primers employed were previously designed and validated to
amplify GAPDH,PECAM1,CDH5, and VWF29. Their nucleotide sequences are
delineated in Supplementary Table 1. All experiments utilized three technical
replicates for each biological sample and included no reverse transcriptase controls.
Data were analysed using Bio-Rad CFX Maestro 1.1 software version 4.1.2433.1219
(Bio-Rad).
The abundance of messenger ribonucleic acid encoding genes of interest in the
putative HAMVECs was compared to that in HUVECs, HCAECs, and HDMVECs
using a two one-sided test for equivalence45. Specifically, equivalence was
established within the significance level αwhen the (1–2α) × 100% confidence
interval of the difference in mean quantification cycles (C
q
) between HAMVECs
and the EC controls was contained within the equivalence margin (± ∂). The
significance level αwas set to 0.05, and the equivalence margin ∂was set to three
standard deviations of the Gaussian distribution of C
q
values amongst the EC
controls about their gene-normalized mean (Supplementary Fig. 3). The EC
controls were evaluated separately but statistically presented as a single population
in order to ascertain the variability of their expression of characteristic endothelial
genes29.GAPDH was used as a loading control.
Immunofluorescence. Cells were rinsed with phosphate-buffered saline (with
calcium chloride and magnesium chloride; PBS+/+), fixed with ice-cold methanol
for 10 min at −20 °C, and blocked with 3% (w/v) BSA in PBS−/−for 30 min prior
to being stained overnight at 4 °C with 5 µg/mL of primary antibodies in the same
blocking solution. The primary antibodies were sourced from Abcam, and included
a mouse anti-human CD31 antibody (catalogue no. ab24590), a rabbit anti-human
VE-cadherin antibody (catalogue no. ab33168), and a mouse anti-human vWF
antibody (catalogue no. ab194405). Cells were then blocked with Normal Serum
Block (BioLegend) for 30 min at 25 °C and stained for 1 hr with 2.5 µg/mL of either
a goat anti-mouse IgG-Alexa Fluor 594 secondary antibody (BioLegend; catalogue
no. 405326) or a goat anti-rabbit IgG-Alexa Fluor 488 secondary antibody (Abcam;
catalogue no. ab150077) diluted in 3% (w/v) BSA in PBS−/−. Cells were then
stained for 5 min with 3 µM 4′,6-diamidino-2-phenylindole (Abcam) in PBS−/−.
Wide-field immunofluorescence microscopy was performed using a Leica DMi8,
operated using the Leica Application Suite X software version 3.5.5.19976
(Leica Microsystems). Image processing was performed using Fiji software version
2.1.0/1.53c46.
Acetylated low-density lipoprotein uptake. AcLDL uptake was assessed as
previously described29. Cells were incubated with 10 µg/mL Alexa Fluor 488-
conjugated AcLDL (Invitrogen) in EGM2 for 4 hr, after which they were fixed with
4% (w/v) paraformaldehyde in PBS−/−for 15 min at 4 °C. Cells were analysed by
flow cytometry.
Angiogenic capacity. The angiogenic capacity of cells was assessed as previously
described47. Briefly, TCPS was coated with 150 µL/cm2of Cultrex PathClear
Basement Membrane Extract (Bio-Techne, Minneapolis, MN, United States). Cells
were then seeded at a density of 45,000 cells/cm2and incubated in EGM2 for 6 hr
before their formation of capillary-like tubules was assessed using phase-contrast
transmission light microscopy (Leica DMIL).
Adipogenic plasticity. The accumulation of lipids by ASCs and HAMVECs was
assessed using a protocol adapted from Kraus et al.48. Cells were seeded onto TCPS
at a density of 4,000 cells/cm2and cultured in EGM2 for 24 hr before the
media was replaced with StemPro Adipogenesis Differentiation Kit (Invitrogen).
The latter was exchanged every other day for 10 days before cells were rinsed with
PBS−/−and fixed with 4% (w/v) paraformaldehyde in PBS−/−for 15 min at room
temperature. Fixed cells were rinsed with 40% (v/v) isopropanol in distilled water
before being stained with 0.2% (w/v) Oil Red O in 40% (v/v) isopropanol in
distilled water for 30 min at room temperature. Stained cells were rinsed with
distilled water before being counterstained with Mayer’s hematoxylin (Abcam;
Cambridge, United Kingdom) for 5 min at room temperature. Cells were rinsed
with distilled water, and representative photomicrographs were acquired using
brightfield transmission light microscopy (Leica DMIL). Oil Red O was then eluted
from the cells by incubating them in 100% isopropanol for 10 min at room tem-
perature, and the absorbance of the eluent at 515 nm was quantified by spectro-
photometry (EnVision 2104 Multilabel Reader operated using EnVision Manager
software version 1.14.3049.528; PerkinElmer, Waltham, MA, United States). The
absorbance of the eluent was compared using a two-way ANOVA with Tukey’s
post-hoc test for multiple comparisons, blocking for donors.
Liquid chromatography–tandem mass spectrometry. Reversed-phase LC–MS/
MS was performed as previously described29. Briefly, cells were lysed and proteins
extracted in 50% (v/v) 2,2,2-trifluoroethanol in PBS–/–supplemented with 100 mM
ammonium bicarbonate. Proteins were reduced with 5 mM dithiothreitol, alkylated
with 15 mM iodoacetamide, and digested with mass spectrometry-grade Trypsin-
Lys C mix (Promega, Madison, WI, United States) following their dilution with
100 mM ammonium bicarbonate and supplementation with 2 mM calcium
chloride. Formic acid was used to quench the digestion, after which the tryptic
peptides were de-salted using OMIX C18 solid-phase extraction tips (Agilent, Santa
Clara, CA, United States). Samples were dried by vacuum centrifugation and
reconstituted in 5% (v/v) formic acid in high-performance liquid chromatography-
grade water.
Tryptic peptides were analysed on an Easy-nLC 1200 (Thermo Fisher Scientific,
Waltham, MA, United States) coupled to a Q Exactive Plus mass spectrometer
(Thermo Fisher Scientific) through a Nanospray Flex Ion Source (Thermo Fisher
Scientific). Tryptic peptides were loaded onto an in-house packed reversed-phase
10-cm, 75 µm internal diameter column (Reprosil-Pur Basic C18, 3 µm, 100 Å; Dr.
Maisch HPLC, Ammerbuch, Baden-Württemberg, Germany), and separated using
a 3-hr acetonitrile linear gradient (2–35% (v/v) in 0.1% (v/v) formic acid in high-
performance liquid chromatography-grade water) at 250 nL/min. All experiments
utilized two technical replicates for each biological sample. Spectra were collected
using a top ten data-dependent acquisition method as previously described29, using
Tune software version 2.8 (Thermo Fisher Scientific) and Xcalibur software version
4.0.27.19 (Thermo Fisher Scientific). Raw files were searched against the UniProt
human proteome database (updated to 2017-07-24) using MaxQuant software
version 1.6.0.1 (Max Planck Institute of Biochemistry, Planegg, Bavaria,
Germany)49, with ‘match between runs’enabled. Cysteine carbamidomethylation
was set as a fixed modification, and methionine oxidation, N-terminal acetylation,
and asparagine or glutamine deamidation were selected as variable modifications.
The false discovery rate (FDR) was set to 1% using a reversed-target decoy
database.
Data visualization and statistical analyses were completed using the Perseus
1.6.1.2 software package (Max Planck Institute of Biochemistry)50. Label-free
quantification values were log
2
-transformed51, and missing values were imputed
from a normal distribution using a downshift of 1.8 and width of 0.3 standard
deviations (Supplementary Data 1 and 2). Unsupervised hierarchical clustering and
associated heat maps of proteins that were identified in ≥2 biological replicates in at
least one group were generated from normalized values across all samples for each
protein. Differentially expressed proteins between groups of cells were defined as
ARTICLE COMMUNICATIONS BIOLOGY | https://doi.org/10.1038/s42003-021-02732-8
12 COMMUNICATIONS BIOLOGY | (2021) 4:1205 | https://doi.org/10.1038/s42003-021-02732-8 | www.nature.com/commsbio
Content courtesy of Springer Nature, terms of use apply. Rights reserved

being either uniquely detected (i.e. detected vs. not detected in ≥2 biological
replicates per group) or quantified in statistically different abundances (i.e. detected
in ≥2 biological replicates in each group, but p< 0.05 or FDR < 0.05). Statistically
overrepresented (p< 0.05) biological pathways (Reactome version 65; released
2019-12-22) among the differentially expressed proteins were identified using the
Fisher’s Exact test with Bonferroni’s correction for multiple testing within
PANTHER version 15.0 released 2020-02-1452, and the Gene Ontological Term
Mapper was used to map differentially expressed proteins to their cellular
components.
Population doubling time. Cells were seeded at a density of 4,000 cells/cm2into
seven TCPS flasks, and every 24 hr cells from one flask were counted using a
hemocytometer. Population doubling times were calculated from the exponential
growth phase using the formula PDT =[Δt× log
10
(2)] ÷ [log
10
(n
f
÷n
i
)], where
PDT represents population doubling time, Δtrepresents the duration of expo-
nential growth, and n
i
and n
f
represent the number of cells at the initiation and
termination of the exponential growth phase, respectively53. The population
doubling times of two populations of cells were compared using two-tailed t-tests,
blocking for donors where appropriate, and those of ≥3 populations were com-
pared using a one-way ANOVA, with Tukey’s post-hoc test for multiple
comparisons.
Population growth rates in co-culture. The effect of seeding purity on the
population growth rates of HAMVECs and ASCs was assessed using an assay
adapted from Gerashchenko (Supplementary Fig. 4)54. HAMVECs and ASCs were
seeded onto TCPS in several pre-defined proportions amounting to a total density
of 4,000 cells/cm2or separately at their respective constituent densities. Cells were
cultured for 4 days, after which the co-cultured cells were stained with 4 µM Cell-
Trace Far Red (Invitrogen) and combined with the mono-cultured controls. The
mixture was then blocked with Human TruStain FcX (BioLegend), stained with
5 µg/mL of mouse anti-human CD31-Alexa Fluor 488 (BioLegend; catalogue no.
303110) for 20 min at 4 °C, and fixed with 4% (w/v) paraformaldehyde in PBS−/−
for 15 min at 4 °C. Cells were analysed by flow cytometry. Population growth rates
in co-culture were calculated using the formulas, R
1
=[Dye+CD31+÷ Dye−
CD31+] × [Dye ÷ Dye−] × 100 and R
2
=[Dye+CD31−÷ Dye−CD31−
] x [Dye+÷
Dye−] × 100, where R
1
and R
2
represent the population growth rates of HAMVECs
and ASCs relative to their mono-cultured controls, respectively. Two-tailed one-
sample t-tests were used to compare the population growth rates in co-culture to
that in mono-culture (i.e. 100%), and a two-way ANOVA with Bonferroni’s post
hoc test for multiple comparisons was used to evaluate the relationship between the
seeding purity and the resultant population growth rates, blocking for donors.
Temporal stability of population purities. The effect of seeding purity on the
temporal composition of cultures was assessed. HAMVECs and ASCs were seeded
onto TCPS in pre-defined proportions (100 or 90% HAMVECs) amounting to a
total density of 4,000 cells/cm2. The purity (% CD31+) of the cultures was assessed
by flow cytometry after 1, 7, 14, and 28 days. The effect of seeding purity on the
purity of the cultures over time was assessed using a two-way ANOVA with
Tukey’s post-hoc test for multiple comparisons, blocking for donors.
Immunomagnetic microparticle size distribution. The size distribution of IMPs
was evaluated with a Multisizer 4e Coulter Counter (Beckman Coulter Life Sci-
ences, Indianapolis, IL, United States), using a 100 µm aperture calibrated with
10 µm beads (Coulter CC Size Standard L10; Beckma n Coulter Life Sciences) or a
20 µm aperture calibrated with 2 µm beads (Coulter CC Size Standard L2;
Beckman Coulter Life Sciences)55. Data was acquired for a modal count of 5,000
using the Multisizer 4e software version 4.03 (Beckman Coulter Life Sciences).
The polydispersity index (PDI) of the IMPs was calculated using the formula
PDI =(standard deviation/mean)2, as previously described55. The size distribu-
tions of the different IMPs were compared using a one-way ANOVA with Tukey’s
post-hoc test for multiple comparisons.
Immunomagnetic microparticle uptake and localization. IMPs were detected on
the basis of their autofluorescence, and in select experiments, a membrane-
impermeable secondary antibody directed against their binding moiety was used to
discriminate between their extracellular and intracellular localization (Supple-
mentary Fig. 5). The autofluorescence of the IMPs and the effect of conjugating the
goat anti-mouse IgG-Alexa Fluor 647 secondary antibody (Cell Signalling Tech-
nology, Danvers, MA, United States; catalogue no. 4410S; staining concentration,
5 µg/mL) on their fluorescence was characterized by spectral scanning confocal
microscopy (Leica SP8 microscope operated by Leica Application Suite X software
version 3.5.5.19976, Leica Microsystems; Advanced Optical Microscopy Facility,
University Health Network), with 20 nm detection bandwidths employed to assess
the emission spectra for each of the 405 nm, 488 nm, 552 nm, and 638 nm exci-
tation wavelengths. 30 IMPs were selected as regions of interest in generating
representative emission spectra. Candidate excitation-emission features of the
IMPs were validated by flow cytometry.
The localization of IMPs in culture was assessed by confocal microscopy. ASCs
were seeded at a concentration of 4,000 cells/cm2onto 35 mm µ-Dishes (ibidi
GmbH, Gräfelfing, Germany) with 400,000 anti-CD31 IMPs/cm2(Dynabeads;
Invitrogen). After 24 hr, cells were rinsed with PBS+/+to remove non-adherent
IMPs, fixed with 4% (w/v) paraformaldehyde in PBS−/−for 15 min at 4 °C, and
permeabilized with 0.1% (v/v) Triton X-100 in PBS−/−for 15 min. F-actin was
stained with Alexa Fluor Plus 647 Phalloidin (Invitrogen), and nuclei were stained
with 3 µM 4′,6-diamidino-2-phenylindole in PBS−/−for 5 min (Abcam). Anti-
CD31 IMPs were detected on the basis of their autofluorescence, and their
localization was assessed by z-stacking confocal microscopy. Images were
processed using Fiji software version 2.1.0/1.53c46.
The prevalence of IMP uptake and their localization in ASCs was assessed by
flow cytometry (Supplementary Fig. 5). Specifically, 105ASCs were exposed to 107
IMPs in suspension for 20 min at 4°C, or in culture in 25cm2TCPS flasks for pre-
defined durations. Cells were then stained with 5 µg/mL of goat anti-mouse IgG-
Alexa Fluor 647 secondary antibody (Cell Signalling Technology; catalogue no.
4410S) for 20 min at 4 °C, and fixed with 4% (w/v) paraformaldehyde in PBS−/−
for 15 min at 4 °C. Cells were analysed by flow cytometry. The prevalence of IMP
localization over time was compared by a two-way ANOVA with Bonferroni’s
post-hoc test for multiple comparisons, blocking for donors; and, the total
prevalence of IMP uptake by ASCs after 20 min in suspension vs. 48hr in culture
was compared using a two-tailed t-test, blocking for donors.
DNA synthesis. The effect of binding and internalizing IMPs on the proliferative
capacity of ASCs was assessed using the Click-iT EdU Alexa Fluor 647 Flow
Cytometry Assay Kit (Invitrogen; Supplementary Fig. 6). ASCs were seeded at a
concentration of 4,000 cells/cm2onto TCPS with 400,000 anti-CD31 IMPs/cm2
(Dynabeads; Invitrogen). After 48 hr in culture, cells were rinsed with PBS+/+to
remove free IMPs and pulsed with 10 µM 5-ethynyl-2´-deoxyuridine (EdU) in
EGM2 for 6 hr. Cells were then resuspended in cell sorting buffer (2 mM EDTA
and 0.1% (w/v) BSA in PBS−/−), and IMP-laden ASCs were magnetically separated
from IMP-free ASCs. Fixation, permeabilization, and staining were performed
using the kit. Cells were analysed by flow cytometry. A two-tailed t-test was used to
compare EdU incorporation between IMP-laden and IMP-free ASCs, blocking for
donors.
Effect of size on immunomagnetic microparticle uptake. The effect of IMP size
on their uptake by ASCs was assessed. Superparamagnetic microparticles ranging
in diameter from 1 to 8 µm (Spherotech, Lake Forest, IL, United States; catalogue
no. SVM-10-10, SVM-40-10, and SVM-80-5) were conjugated to mouse anti-
human CD31 antibodies (Miltenyi Biotec; catalogue no. 130-119-893) via biotin-
streptavidin interactions (5 µg of antibody per 107particles). Their uptake by ASCs
was compared to that of the commercial anti-CD31 IMPs (Dynabeads CD31
Endothelial Cell; Invitrogen) and the anti-CD31 cIMPs. The size distributions of
these five distinct anti-CD31 IMPs was assessed using a Coulter counter.
The binding and internalization of the anti-CD31 IMPs was assessed by flow
cytometry (Supplementary Fig. 9b and 9c). Specifically, 105ASCs were exposed to
107IMPs in suspension for 20 min at 4 °C, or in culture in 25 cm2TCPS flasks for
pre-defined durations. Cells were then resuspended in cell sorting buffer (2 mM
EDTA and 0.1% (w/v) BSA in PBS−/−), and IMP-laden ASCs were magnetically
separated from IMP-free ASCs. IMP+ASCs were stained with 4 µM CellTrace Far
Red (Invitrogen) before being re-combined with the IMP−ASCs. The mixture was
then stained with 5 µg/mL of goat anti-mouse IgG-Alexa Fluor 488 secondary
antibody (Invitrogen; catalogue no. A11001) for 20 min at 4 °C, and fixed with
4% (w/v) paraformaldehyde in PBS−/−for 15 min at 4 °C. Cells were analysed by
flow cytometry. The effect of IMP size on their uptake by ASCs over time was
assessed using a two-way ANOVA with Tukey’s post-hoc test for multiple
comparisons, blocking for donors.
Enrichment efficacies of alternative MACS strategies. The effects of alternative
target antigens, sizes, and exposures of IMPs on the enrichment efficacy of the
MACS procedure were investigated using an in vitro model of contaminated pri-
mary cultures. HAMVECs and ASCs were seeded onto TCPS at a density of
4,000 cells/cm2in a 9:1 proportion with or without 400,000 IMPs/cm2to recapi-
tulate their potential exclusion from primary cultures. Their enrichment was
performed using MACS after 4 days, and their purities (% CD31+) were evaluated
after a total of 7 days by flow cytometry. A two-way ANOVA with Tukey’s post-
hoc test for multiple comparisons was used to compare the culture purities of the
different MACS strategies, blocking for donors. The effects of target antigen and
IMP size/exposure were assessed using a one-way ANOVA with Tukey’s post-hoc
test for multiple comparisons; pooling replicates where applicable.
Statistics and reproducibility. Experiments were performed three times using
cells derived from three different donors (n=3 biologically independent samples),
unless indicated otherwise. Other than the proteomics data that was analysed using
the Perseus 1.6.1.2 software package (Max Planck Institute of Biochemistry)50,
statistical analyses were performed using Prism 8 software version 8.4.3 (GraphPad
Software, San Diego, CA, United States). Where applicable, normality of data was
assessed using quantile-quantile plots and the Shapiro–Wilk test. Unless stated
otherwise, p< 0.05 was accepted as statistically significant and values are repre-
sented as mean ± standard deviation.
COMMUNICATIONS BIOLOGY | https://doi.org/10.1038/s42003-021-02732-8 ARTICLE
COMMUNICATIONS BIOLOGY | (2021) 4:1205 | https://doi.org/10.1038/s42003-021-02732-8 | www.nature.com/commsbio 13
Content courtesy of Springer Nature, terms of use apply. Rights reserved

Reporting summary. Further information on research design is available in the Nature
Research Reporting Summary linked to this article.
Data availability
Raw data and search results from the LC–MS/MS are available from the MassIVE
repository (accession no. MSV000086982), and the corresponding tabulated datasets are
supplied in Supplementary Data 1 and 2. Source data underlying the graphs and charts
presented in the main figures are provided in Supplementary Data 3. All other data are
available from the corresponding author upon reasonable request.
Received: 23 April 2021; Accepted: 27 September 2021;
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Acknowledgements
The authors thank Kate Butler for her assistance in facilitating the provision of tissue for
this research. This work was funded by the Canadian Institutes for Health Research
(grant no. 230762 & 426275). J.A.A. was supported by a Post Graduate Scholarship—
Doctoral (PGS-D3) from the Natural Sciences and Engineering Research Council of
Canada (NSERC), an Ontario Graduate Scholarship, and an Education Fund award from
the Ted Rogers Centre for Heart Research (TRCHR). V.M. was supported by an
Undergraduate Student Research Award from NSERC and a Ted Rogers Scholar Award
from TRCHR.
Author contributions
J.A.A., C.A.S. and J.P.S. conceived the study. J.A.A. designed the study. J.A.A., V.M. and
M.J.M. collected and analysed data. A.O.G., S.O.P.H. and J.P.S. provided material for the
study. J.A.A. wrote the article. All authors reviewed and approved the article.
Competing interests
The authors declare no competing interests.
Additional information
Supplementary information The online version contains supplementary material
available at https://doi.org/10.1038/s42003-021-02732-8.
Correspondence and requests for materials should be addressed to J. Paul Santerre.
Peer review information Communications Biology thanks Emad Moeendarbary and the
other anonymous reviewers for their contribution to the peer review of this work.
Primary Handling Editors: Marco Fritzsche and Eve Rogers. Peer reviewer reports are
available.
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COMMUNICATIONS BIOLOGY | https://doi.org/10.1038/s42003-021-02732-8 ARTICLE
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