1. Introduction
The notion of the mammalian adult heart as a terminally differentiated organ has been challenged during the last 20 years [
1]. Cardiomyocyte (CM) proliferation has been demonstrated in young individuals (<20 years old) [
2] and upregulated in some pathological conditions, like myocardial infarction (MI) [
3]. Despite different hypotheses for cell turnover being explored in cardiac tissue, the source(s) of new cells in the adult mammalian heart remain enigmatic (reviewed by [
4]). Cardiomyocyte turnover has been proposed, being sustained by the de-differentiation/proliferation/re-differentiation of certain mature mouse CM; however, this proposal has demonstrated a very low frequency (<1% of annual turnover) and still awaits more robust empirical verification [
5].
Tissue homeostasis, repair, and damage response mainly relies on the regulated activity of scarce populations of tissue-specific adult stem cells (ASCs)/progenitors. Classical studies on ASCs have relied on the use of supposed specific ASC markers and tracing of their progeny. In tissues with a high cell turnover, these ASC populations are clearly defined (reviewed by [
6,
7,
8]). Regarding the adult heart, several markers have been proposed for the identification/isolation of cardiac resident stem/progenitor cells (multipotent, CSC) [
9]. Among them,
c-Kit+ cardiac cells were the first and the most intensively studied candidate population [
10,
11] as a potential resource for cardiovascular therapy. However, later findings from basic and preclinical research, together with the failure of several clinical trial evaluations, have fueled a long and bitter debate over the relevance of
c-Kit+ CSC [
12]. Other progenitor-like cell populations have been described, most of these proposals being based on cell surface proteins that are also expressed on other ASCs or markers associated with general progenitor functions, such as
Sca1,
Abcg2, or
Isl1 (reviewed in [
13]). As an independent strategy to define elusive CSC, expression of
Bmi1 as the most representative marker of mouse adult stem cell compartments (reviewed by [
14]) was evaluated in the adult mouse heart.
BMI1 is a member of the Polycomb Repressive Complexes 1 (PRC1), a well-recognized transcriptional suppressor with the ability to drive self-renewal and proliferation of many tissue-specific stem cells [
15]. Using appropriated mouse models, we confirmed the existence of a non-cardiomyocyte
Bmi1+ population; around 75,000
Bmi1+ cells were estimated in the mouse adult heart [
15].
Bmi1+DR cells contain two major, and mutually exclusive, subpopulations: PDGFRα+ CD31- and CD31+ PDGFRα- populations. In previous studies, we have demonstrated the in vitro and in vivo multilineage cardiac differentiation capacity of
Bmi1+DR cells, showing their contribution to the basal “wear and tear” of cardiac endothelial cell (EC), vascular smooth cell (SMC), and CM lineages, with a substantial endothelial bias both in homeostasis and in response to acute myocardial infarct (AMI) as well as other forms of cardiac damage [
15,
16,
17,
18,
19]. Transcriptional analysis demonstrated that
Bmi1 expression is linked to a mixture of endothelial- and mesenchymal-related non-myocyte
Sca1+ cells in the adult mouse heart, although the level of expression of endothelial genes was significantly lower in
Bmi1+ cells when compared with mature endothelial cells [
17,
18,
19].
Regarding tissue repair, solid evidence has demonstrated that cardiac
Bmi1+ progenitors are quite resistant to several forms of damage, like AMI, instead becoming proliferatively activated, with a net increase in their cell numbers during the first few days [
16,
17,
18]. Moreover, their progeny showed an enhanced contribution to the main de novo cell lineages, including CM, when compared with cells from age-paired non-infarcted adult hearts [
18,
20]. Due to this capacity, this population is called
Bmi1+DR cells for
Bmi1+ Damage-Responsive [
19]. Redox status influences
Bmi1+DR cells response and highlights redox-mediated BMI1 regulation, with implications for the maintenance of cellular identity in vivo [
17]. It was estimated that at 4 months post-AMI, the
Bmi1+DR population contributes up to 20% of total endothelial cells in the infarcted heart [
18]. The relevant in vivo physiological role of the
Bmi1+DR population has been validated by a genetic ablation model; when
Bmi1+DR cell ablation was coupled to AMI, animals manifested signs of cardiac dysfunction, affecting the survival. Perimortem analysis revealed a dilated cardiomyopathy-like phenotype with a significant deficit in the angiogenic response to AMI as the most probable cause of death [
18].
The ASC niche, defined as the local (micro)environment surrounding a stem cell-containing population, is now recognized as the functional unit for ASC maintenance and regulation [
21]. Stem cell niches are dynamic functional domains rich in specialized cells that influence, condition, and coordinate ASC behavior to govern tissue homeostasis under physiological conditions, but in certain contexts, the stem cell niche can be corrupted, as in some cancers and chronic pathologies [
22,
23,
24]. ASC niche might function both through direct cell–cell contact and by releasing soluble factors. However, recent studies on the long-term lympho-hematopoietic stem cell (HSC) niche, undoubtedly the best-characterized model of the ASC niches, have uncovered new layers of regulatory complexity. For instance, HSC progenies themselves regulate HSC behavior, lineage-biased differentiation of HSCs is coordinated by distinct niches, and HSCs can remodel their own perivascular niche [
25,
26,
27]. Currently, based on niche composition, this well-described perivascular niche has been defined in other ASCs (e.g., neural stem cells, cancer stem cells) with a similar level of complexity. In any case, scarce research has been invested in the characterization of adult heart niches.
Aiming to identify and define the
Bmi1+DR cell niche(s), we analyzed mouse adult heart sections to find that the majority of
Bmi1+DR cells were located in the left ventricle (≈70%) in a gradient-like distribution around the cardiac vasculature, and preferentially close to small vessels [
19]. Interestingly, in the adolescent heart and earlier ages,
Bmi1+DR cells display an almost random distribution, suggesting that perivascular confinement becomes relevant in an age-dependent manner, with the progressive increasing of oxidative stress. Results confirmed that
Bmi1+DR cells are sheltered in low-ROS perivascular domains, allowing us to propose that these structures form part of the perivascular niche for the
Bmi1+DR population in the adult heart [
19] (reviewed by [
13]). Furthermore, in vivo evaluation of
Bmi1+DR cells proliferative status relative to cardiac vasculature demonstrated that only a small percentage of these cells (≈10% of total) were proliferating. Equivalent analysis on infarcted hearts showed a strong distortion of the spatial distribution of
Bmi1+DR cells in relation to cardiac vessels [
19]. Altogether, these results showed that non-proliferating
Bmi1+DR (quiescence-like) cells are preferentially located in strict areas close to the endothelium in homeostasis, strongly suggesting functional interactions. Preliminary evaluation [
19] confirmed that, specifically, the co-culture of
Bmi1+DR with endothelial cells decreases both of their proliferation rates, incrementing the expression of
Bmi1 as expected [
28,
29]. In direct contact, endothelial cells also promoted a reduction in total ROS, which was concomitant with a decrease in total mitochondrial mass, a hallmark of
Bmi1+DR [
19].
In conclusion, as in other ASC models, a perivascular niche for Bmi1+DR cells is envisioned. Better comprehension of the regulation of the cardiac niche(s) would be key for resolving uncertainties about the involvement of cardiac progenitor cells/stem cells in heart homeostasis and damage repair, and for demonstrating whether the low margin of heart turnover is relevant for healthy aging or counteracting some pathological scenarios. Then, in this study, we generated and characterized an in vitro Bmi1+DR immortalized cell model in an attempt to overcome some of the technical limitations associated with work involving such a scarce population of primary cells. Additionally, we applied this model to unravel some molecular mechanism(s) that could define the nature of this niche-like relationship among Bmi1+DR cells and the cardiac endothelium.
3. Discussion
The niche relationship among different ASC populations and the endothelium has been widely defined in several organs/compartments, for example, in neural stem cells (NSCs) [
45,
46,
47], bone marrow (BM)-derived cells [
48,
49], and skeletal muscle stem cells (MuSCs) [
50,
51]. In our previous studies, we confirmed the spatial relationship observed between
Bmi1+DR cells and the cardiac endothelium [
19]. Then, in an attempt to try to understand the role of the
Bmi1+DR population in cardiac turnover and response to damage, we consider a key point to unravel their potential niche-like relationship with cardiac endothelium.
This niche relationship is envisioned as critical for the maintenance and fate of
Bmi1+DR cells. Previously, it was confirmed that after AMI (5–10 d),
Bmi1+DR cells are not apparently damaged; they were even proliferatively activated [
18], moderately increasing their numbers, and as the main consequence, long after AMI (4 months), a substantial increase in the number of mature cells is demonstrated (
Figure S1B). Here, we confirmed that a similar behavior after Pq treatment (single dose, 5 d post-Tx) is inducible in a Bmi1-Tomato mouse model (
Figure 1A(II)); monitoring of heart cryosections of the
Tomato+ (
Bmi1+) signal, and cell numbers, relative to the total (DAPI+) cells, also showed a moderate expansion (
Figure 1E). Similarly, it is established in mammalian arteries that smooth muscle cells (SMCs), a highly resting population in human arteries, only contribute efficiently to repair when drastic damage is inflicted [
44].
Cardiac endothelium, as the proposed niche for the
Bmi1+DR population, might control its cell distribution, differentiation potential, and proliferative status. A previous study showed that
Bmi1+DR cells display a perivascular gradient-like cell distribution in the adult mouse heart, with only a small percentage of these cells (≈10% of total
Bmi1+ cells) being in a proliferative state [
19]. Moreover, Herrero at al. showed how oxidative damage conditions modified
Bmi1 activity in vivo by derepressing canonical target genes in favor of their antioxidant and anticlastogenic functions to trigger ROS-associated differentiation of this cardiac progenitor population, pointing out that the differentiation potential of
Bmi1+DR cells is clearly mainly controlled by oxidative stress [
17]. In addition, we found that, specifically, only perivascular areas with very low ROS levels coincided with the localization of the majority of
Bmi1+DR cells in vivo, which is very similar to those described in other ASC compartments [
29], but this specific distribution was distorted when general low-ROS conditions were applied by genetically decreasing ROS levels in Bmi1-Tomato mice using G6PDTg, which confirmed the importance of perivascular ROS levels [
19]. On the contrary, here we showed that when general high-ROS conditions were applied with Pq treatment,
Bmi1+DR cell preference to be close to the endothelium increases, showing a tighter relationship with the endothelium in response to damage (
Figure 1F). All of these data reinforce the vascular niche hypothesis for
Bmi1+DR cells, with oxidative stress levels being a major regulator (
Figure 1G).
In order to achieve a deeper analysis of this niche-like relationship of
Bmi1+DR cells with cardiac endothelium, and to unravel the regulatory mechanisms, we decided to carry out an analysis in cultures of
Bmi1+DR and endothelial cells, as conducted previously [
19]. However, due to the scarcity of
Bmi1+DR cells, we faced significant experimental limitations, as we were restricted to working with primary cultures, in addition to the great number of mouse models needed to try to understand the physiology of this progenitor population. Then, aiming to facilitate the dissection of the plausible mechanisms that could play a relevant role involving endothelial cells and
Bmi1+DR cells, as well as to reduce the needs for animal models, we tried to develop a reversible immortalization procedure for
Bmi1+DR cells.
After revision and evaluation of the previous literature, we decided to use the expression of the SV40-T that was successfully used with similar cell types [
31,
32]. Other strategies involved the overexpression of
Bmi1 and
TerT, which were both discarded because of the central role of
Bmi1 expression in
Bmi1+DR cells. In addition, it was demonstrated that in primary cardiomyocytes, SV40-T is a superior immortalization agent when compared with
Bmi1 and
TerT [
36]. We aimed to immortalize the
Bmi1+DR population, isolated from the inducible Bmi1-YFP mouse model (
Bmi1+/+) (
Figure 1A(I)) [
15,
52]. This population was successfully immortalized, generating the
Bmi1+DR
IMM pool, which showed a clear exponential and maintained (more than 40 passages) proliferation (
Figure 2C). The
Bmi1+DR
IMM population was characterized, demonstrating a similar expression of relevant cell surface makers, global expression profiles, and biological responses to several signaling molecules, previously tested to define
Bmi1+DR cells [
15,
16,
17,
18,
19] (
Figure 2F–H). As previously described,
Bmi1+DR cells were initially considered as a mixed population of mutually exclusive PDGFRα+ and CD31+ cells. Although flow cytometry of
Bmi1+DR
IMM showed negative results for CD31 (
Figure 2F) expression, we considered that a strong post-transcriptional effect could be involved or the PDFGRα+ subpopulation could have a higher proliferating rate [
18]. Then,
Bmi1+DR
IMM should represent the
Bmi1+DR/PDGFRα+ cells.
Nonetheless, our first attempt was to develop a reversible immortalization procedure where, upon the transient expression of Cre-Recombinase, the LoxP flanking sequences would recombine and delete the immortalization cassette; furthermore, this construct also expresses TK to eliminate cells that would not be properly engineered. Unfortunately, although reversibility of SV40-T immortalization has been described [
23,
53,
54], the procedure on
Bmi1+DR
IMM provoked a sudden proliferation block and the induction of senescence (
Figure 3). This phenotype has been also found in other cell types, such as human olfactory ensheathing glia [
35]. This result could not be rescued by modifications of the culture medium nor by the addition of senescence inhibitors (
Figure S3). In conclusion, although we could not fully use the
Bmi1+DR
IMM platform as planned, the
Bmi1+DR
IMM population was confirmed as an incredibly useful tool to study these cardiac progenitor cells, postulating this immortalization procedure to be applied for the study of other complex adult progenitor populations. Altogether, the
Bmi1+DR
IMM population seems to be quite a reasonable model for the study of
Bmi1+DR cells endothelial niche.
The niche concept was first proposed by R. Schofield for HSCs and referred to the surrounding supporting cells and the soluble factors that influence HSC behavior [
55]. Currently, the niche is generally considered the real functional unit in most ASC compartments, being critically responsible for tissue or organ homeostasis, damage response (regeneration), and repair [
6,
7,
56]. Among all ASC models, HSCs [
57,
58], NSCs [
45,
59], and MuSCs [
50,
51] are, perhaps, the best-known compartments. Many data postulate that MuSCs [
60] and
Bmi1+DR cells [
19] showed a similar interaction with their corresponding microvasculature, mainly controlling them by redox regulation [
13,
17,
18,
19,
61]. One of the main goals of the potential cardiac vascular niche regulation is to reduce the impact of the progressive oxidative stress in the progenitor populations during adulthood, aging, or cardiac damage conditions. However, to our knowledge, no study has addressed the plausible direct impact of experimental oxidative stress levels (acute or chronic) on the biology of MuSCs and their niche, as well as the main cell–cell interactions contributing to the preservation of MuSCs and counteracting oxidative stress; research has been concentrated in ischemia- and reperfusion-derived damage aging [
62], as well as in other experimental dedicated models. Then, in the cardiac tissue, we asked whether vasculature could protect
Bmi1+DR cells against substantial ROS levels.
In this regard, we first analyzed
Bmi1+DR
IMM cells response to oxidative damage by in vitro Pq treatment and demonstrated quite a similar response to the non-immortalized population (
Figure 4A,B), as well as to the analysis of the whole heart [
39]. Next, we modelled in vitro the “minimal” cardiac endothelial niche, co-culturing
Bmi1+DR
IMM cells (or, when indicated, primary
Bmi1+DR cells) with the 1g11 endothelial cell line or primary cardiac endothelial cells (pCECs), and evaluated the functional consequences of short acute oxidative stress conditions (Pq, 12 h) compared with individually cultured
Bmi1+DR cells (mono-culture) and other majoritarian cell types in the cardiac tissue. Results clearly indicate that co-culture of
Bmi1+DR
IMM with the 1g11 endothelial cell line and pCECs, but not with embryonic fibroblasts (MEFs) or the mouse cardiomyocyte-like cell line HL-1, promotes a protective effect for resistance to medium–high Pq concentrations (5 or 8 mM) (
Figure 4F–I). The effect was dose-dependent and specific, as HL-1 cells were not protected by endothelial co-culture (
Figure S4C), and the effect was unidirectional, as
Bmi1+DR
IMM cells did not exert any protective effect on the other cell populations (
Figure S4D–F). Therefore, it can be concluded that endothelial cells seem to play a notable role in preserving
Bmi1+DR maintenance from oxidative stress. This consideration must be related to the high in vivo resistance of
Bmi1+DR cells to several forms of damage, including AMI, irradiation, mitomycin [
17], and Pq. This protective role of niche endothelial cells in stem cell populations was first described in the lymphohematopoietic system after radiation [
63] and in cardiac resident populations. Moreover, it was previously described that the cardiac Sca1+ CD31- subpopulation protects cardiomyocytes against different forms of damage, included AMI, being mediated by MCP-1 [
64] and potentiated by miR-133a [
65].
On the other hand, a central and solid result concerned the substantial reduction in
Bmi1+DR intracellular ROS by direct co-culture with pCECs [
19], so we evaluated the regulation of the main mechanisms that could contribute, including autophagy and metabolism shift, as revealed for skeletal muscle [
66]. Autophagy is key in preventing stresses as one of the major quality control guardians in the cell; the autophagy pathways acquire physiological relevance even under basal, non-stressful conditions, being especially relevant for the maintenance of stem cell self-renewal potential [
67,
68], cellular differentiation, and plasticity [
69]. Tissues that are mainly composed of post-mitotic/quiescent cells exhibit higher sensitivity to loss of autophagy competence. For example, in the skeletal muscle, MuSCs display a continuous basal level of autophagy critical for their stemness and maintenance capacity, showing that physiological decline in autophagy in old satellite cells or its genetic impairment in young cells results in toxic cellular waste accumulation and progression towards senescence [
70]. In this context, we aimed to evaluate whether primary endothelial cells (pCECs) could modulate autophagy in
Bmi1+DR cells, after direct contact co-culture of these cells. Results indicated that, unexpectedly, autophagy was not enhanced but, on the contrary, total and non-canonical autophagy flux was moderately reduced by co-culture with pCECs. This result was further confirmed by RT-qPCR analysis of the panel of genes relevant for autophagy; none of them were enhanced but, on the contrary,
Bnip3L and Bnip3, critically involved in mitophagy (selective autophagic degradation of mitochondria) [
44,
71], appeared quite significantly reduced only in direct contact co-cultures (65 and 80% reduction, respectively). According to this, in the context of aging, basal autophagy was found to be reduced in a subset of younger HSCs compared to their older counterparts [
72]. In this way, while mitophagy might be critical for clearing metabolically active mitochondria to maintain quiescence in some stem cell populations [
72], the role of mitochondria is key in NSCs, as mitochondria regulate self-renewal by maintaining low levels of ROS [
73]. Therefore, although we did not find direct evidence that pCECs, in the basal stage, promote autophagy in
Bmi1+DR
IMM cells by cell–cell contact, it is possible that the reduction in non-canonical autophagy and mitophagy-associated gene expression might be associated with the regulation of ROS levels by the endothelial niche. Moreover, some critical signals (molecules or other cell types) could be lost from this minimal vascular niche.
Nonetheless, another critical mechanism involved in intracellular ROS control among ASC populations is the regulation of their metabolic activity. Indeed, their relationship with the vascular niches has been also related with metabolic control. For example, HSCs in homeostasis reside close to their vascular niche, which promotes an enhanced glycolytic state, reducing ROS production [
74]. Equally, the opposite effect has been observed in studies performed in zebrafish, with an enhanced oxidative metabolism in endothelial cells as an initiator of the revascularization process after cardiac damage [
75]. Then, we decided to study the effects on
Bmi1-DR
IMM cell metabolism of pCEC direct co-culture. The analysis of the two main metabolic pathways did not render important differences; the co-culture with pCECs compared to individually cultured
Bmi1+DR cells rendered a moderately higher oxygen consumption rate index (OCRI, proportional to mitochondrial respiration), particularly for the maximal respiratory rate, but a similar basal respiration and no differences in extracellular acidification rate (ECAR, proportional to glycolysis) were found. However, in a second analysis by RT-qPCR, we observed that co-culture with pCECs reduced
Apelin expression in
Bmi1+DR
IMM cells, a critical metabolic mediator that activates import and consumption of glucose and fatty acids [
76],
Ppargc1a, a master regulator of oxidative metabolism and some targets, and
Atp5j [
77]. These modifications of expression profile would suggest that co-culture with pCECs promotes a metabolic slow-down, as in other stem cell compartments [
41]. Accordingly, quiescent hematopoietic stem cells exhibit low oxidative phosphorylation levels, switching to a high-oxidative-phosphorylation metabolic state only after their activation. Globally, further studies are required to determine if autophagy, or even mitophagy, and metabolic glycolytic state play a role in modulating the switch in ROS level dynamics between quiescent undifferentiated and differentiated
Bmi1+DR cells by their endothelial niche.
While our current study demonstrates that the
Bmi1+DR
IMM pool not only maintains long-term cell proliferation capacity but also retains their native
Bmi1+DR cell counter-part characteristics, some differences and limitations should be pointed out. In relation to the immortalization process, the reversibility of SV40-T immortalization on
Bmi1+DR
IMM cells was unsuccessfully achieved due to the fulminant senescent phenotype provoked; this anticipates limitations of their clean usage for certain applications. Obviously, we were interested in a well-controlled immortalization for expansion, followed by dis-immortalization prior to functional evaluations. The global phenotype seems to be related to the selection of the immortalizing function, because a previous publication using SV40-T found a similar picture, demonstrating that cell proliferation rate significantly decreased in selected clones of cardiac progenitor cells upon SV40-T removal [
53]. In spite of that, all of the discussed evidence indicates that the
Bmi1+DR
IMM pool reasonably resembles the main characteristics of primary
Bmi1+DR cells, including some biological and oxidative responses [
19], but reflecting also clear differences (
Figure 2F–H); all data suggest that
Bmi1+DR
IMM cells might better represent a
Bmi1+DR/PDGFRα+ subpopulation. Overall, the
Bmi1+DR
IMM pool was proven to be a useful tool, allowing us to recreate the minimal vascular niche using co-cultures and continue its definition. Further work with the
Bmi1+DR
IMM pool has also demonstrated that they can be used for colony forming assays with quite similar results to primary
Bmi1+DR cells, and even in in vitro forced differentiation assays. We are convinced that further work with
Bmi1+DR
IMM cells will help to expand complexity in the cardiac minimal vascular niche similarly to the composition and regulation of the skeletal muscle niche, as the closest reference.
4. Materials and Methods
4.1. Transgenic Mice and Tamoxifen Administration
Transgenic mice used in this study,
Bmi1CreERT/+-
Rosa26YFP/+, Bmi1CreERT/+-
Rosa26TdTomato/+ and
Bmi1GFP/+ (all from The Jackson Laboratory), were maintained on the C57BL/6 background, as previously required [
15,
16,
17,
18,
19]. All animal strains used were adult mice (8–12 weeks old); as previously indicated in previous studies and detailed in
Supplementary Materials S2.1, they are used with the corresponding administrative and ethical authorizations.
For Tamoxifen (Tx) administration, Tx (Sigma-Aldrich Inc., St. Louis, MI, USA, T5648) was dissolved in corn oil (Sigma-Aldrich Inc., St. Louis, MI, USA, C8267) and intraperitoneally (i.p.) injected (225 μg/g body weight) in Bmi1CreERT/+-Rosa26YFP/+ or Bmi1CreERT/+-Rosa26TdTomato/+ animals every 24 h for 3 days. The animals were used, fundamentally, for the different experiments 5 days after finishing the induction.
4.2. Immunofluorescence of Cardiac Tissue and Image Analysis
The extraction of cardiac tissue was performed 5 days after the last dose of Tx, or 48 h after paraquat (Pq) administration. Once the animals were anesthetized and sacrificed, the heart was perfused with 1X PBS through the hepatic vein to clean the remains of blood from the ventricular and atrial cavities. After that, the heart was kept rotating for 12 h at 4 °C in a solution of 4% paraformaldehyde (PFA; TED PELLA, Redding, CA, USA, 18505) for fixation. Afterwards, the heart was kept in increasing sucrose solutions in a gradient of 15 to 30% concentration for dehydration. This allows final inclusion in OCT (Sakura Finetek Spain S.L., Barcelona, Spain, 25608-930) of the heart. Using a microtome, histological sections of 6–8 µm thickness were obtained from the ventricular zone of each heart for subsequent analysis by immunofluorescence.
The histological heart sections were treated for 1 h at room temperature (RT) with 1X PBS + 3% bovine serum albumin (BSA; Sigma-Aldrich Inc., St. Louis, MI, USA, A7906). The permeabilization of the membrane was performed with the detergent Triton X-100 dissolved at 0.5% in 1X PBS, incubating the sections at RT for 20 min. After several washes with 1X PBS + 1% BSA, histological sections were incubated with blocking solution 1X PBS + 5% BSA for 2 h at RT. After washing again with 1X PBS + 1% BSA, sections were incubated overnight at 4 °C with the corresponding primary antibodies, Rabbit anti-αSMA (Abcam, Cambridge, England, ab5664) and Rat anti-Sca1 (RyD systems, Minneapolis, MN, USA, MAB1226), diluted to 1:50 and 1:100, respectively, in a solution of 1X PBS + 1% BSA + 0.1% Triton X-100). The next day, sections were washed with 1X PBS + 1% BSA and subsequently incubated for 1 h at RT with the corresponding secondary antibodies (anti-Rabbit 647 nm and anti-Rat 488 nm (Jackson, Bar Harbor, ME, USA, 111-176-104) prepared at 1:500 in the same solution of the primary antibody. After several washes with 1X PBS + 1% BSA, sections were incubated with DAPI (Sigma-Aldrich Inc., St. Louis, MI, USA, D9542) diluted in 1X PBS at a concentration of 1:500 for 20 min at RT. Finally, sections were set up with ProLong Antifade Mountant (Invitrogen, Madrid, Spain, P36930). The resulting immunofluorescence of the cardiac tissue was analyzed by imaging with the Leica SP5 Microfluor microscope and their subsequent processing and analysis with the Image J program version FIJI (National Institute of Health, Belthesda, MD, USA).
4.3. Isolation and Culture of Adult Mouse Non-Myocyte Bmi1+DR Cells
Primary non-myocyte cells and cardiomyocytes were obtained by the Langendorff method using retrograde perfusion through the aorta. The heart was removed rapidly and retrograde-perfused under constant pressure (60 mmHg; 37 °C, 8 min) in Ca
2+-free buffer (113 mM NaCl, 4.7 mM KCl, 1.2 mM MgSO
4, 5.5 mM glucose, 0.6 mM KH
2PO
4, 0.6 mM Na
2HPO
4, 12 mM NaHCO
3, 10 mM KHCO
3, 10 mM Hepes, 10 mM 2,3- butanedione monoxime, and 30 mM taurine). Digestion was initiated by adding a mixture of recombinant enzymes (0.2 mg/mL Liberase Blendzyme (Roche, Madrid, Spain, 05401127001), 0.14 mg/mL trypsin (ThermoFisher, Waltham, MA, USA, 15090046), and 12.5 μM CaCl
2) to the perfusion solution. When the heart became swollen (10 min), it was removed and gently teased into small pieces with fine forceps in the same enzyme solution. Heart tissue was further dissociated mechanically using 2, 1.5, and 1 mm diameter pipettes until all large heart tissue pieces were dispersed. The digestion buffer was neutralized with stopping buffer (10% fetal bovine serum (FBS; Capricorn Scientific, Ebsdorfergrund, Germany, FBS-12A) and 12.5 μM CaCl
2). Cardiomyocytes were pelleted by gravity in a two-phase decantation process (45 and 30 min, respectively), and the supernatant was used as a source of non-myocyte cardiac cells [
14].
Primary Bmi1+DR cells were isolated from Bmi1CreERT/+-Rosa26YFP/+ (Bmi1+DR YFP+ cells), Bmi1CreERT/+-Rosa26Tomato/+ (Bmi1+DR Tomato+ cells), and Bmi1GFP/+ (Bmi1+DR GFP+ cells) mice by cell sorting with the corresponding reporter after Langendorff digestion and expanded in Iscove’s modified Dulbecco’s medium (IMDM; ThermoFisher, Waltham, MA, USA,12440-053) supplemented with 10% FBS, 100 IU/mL penicillin (Invitrogen, Madrid, Spain), 100 mg/mL streptomycin (Invitrogen, Madrid, Spain), 103 units ESGRO-LIF (Millipore, Burlington, MA, USA, ESG1107), 20 ng/mL FGF (Fibroblast Growth Factor; Peprotech,100-18B), 10 ng/mL EGF (epidermal growth factor; Peprotech, AF-100-15), and 100 μg/mL Normocin (InvivoGen, San Diego, CA, USA, ant-nr-1). Bmi1+DR cells were cultured under hypoxic conditions (37 °C, 3% O2, 5% CO2) and culture plates previously treated with 0.1% gelatin.
4.4. Isolation and Culture of Adult Mouse Primary Cardiac Endothelial Cells
For the isolation of primary cardiac endothelial cells (pCECs), wild-type (WT) mice that did not include any genetic modification were used. After euthanizing the animals, the hearts were perfused with 1X PBS through the vena cava to eliminate circulating hematopoietic cells in the chambers of the heart that could interfere with the subsequent extraction process. The heart was removed and was mechanically disintegrated with the help of a scalpel. Once the heart was disintegrated into the smallest fragments possible, we proceeded with enzymatic digestion using DMEM medium supplemented with Collagenase (Sigma-Aldrich Inc., St. Louis, MI, USA, C5138) and Dispase II (Hoffmann-La Roche, Basel, Switzerland, 04 942 076 001), both at a concentration of 1 mg/mL, at 37 °C for 45 min under stirring. Homogenization of the resulting solution was performed by passing it through a sterile 18 G needle and a 70 µm sterile filter (Sigma-Aldrich Inc., St. Louis, MI, USA, 352350) to eliminate possible large fragments not digested correctly. Isolation medium (DMEM supplemented with 20% FBS, 100 U/mL penicillin, and100 µg/mL streptomycin) was added and the resulting cell suspension was centrifuged at 400 g for 5 min. After washing with 1X PBS + 0.5% BSA, cells were centrifuged at 300× g for 10 min.
pCECs were isolated from the obtained pellet by magnetic separation using the MACS Neonatal Cardiac Endothelial Cell kit Isolation Kit (MACS Miltenyi Biotec, Bergisch Gladbach, Germany, 130-104-183) and expanded in VascuLife VEGF Endothelial Medium Complete Kit (Lifeline Cell Technology, San Diego, CA, USA, LL-0003). pCECs were cultured under normoxic conditions (21% O2, 5% CO2, 37 °C) in plates previously treated with 1% gelatin and supplemented with 100 μg/mL fibronectin (Sigma-Aldrich Inc., St. Louis, MI, USA, F1141); cells were used for the experiments at passage ≤4–5.
4.5. Culture Conditions for Other Cell Lines
4.6. Immortalization/Dis-Immortalization of Bmi1+DR Cells
4.6.1. Immortalization, Transduction of Bmi1+DR Cells with the Lentiviral LoxP-SV40 T-Large–TK-LoxP Vector, and Further Expansion/Confirmations
To obtain the immortalized pool of primary
Bmi1+DR, cells were isolated and sorted from
Bmi1CreERT/+-
Rosa26YFP/+ (
Bmi1+DR YFP+ cells) or
Bmi1GFP/+ (
Bmi1+DR GFP+ cells) mice and transduced with the immortalization lentiviral vector (pLOX-Ttag-iresTK; Addgene, Watertown, MA, USA, 12246); the composition of the vector is depicted in
Figure 2A. The pLOX-Ttag-iresTK vector was produced by the viral vector production unit at the National Center for Cardiovascular Research (CNIC) (Madrid, Spain), and it is a 3rd-generation lentiviral vector, in terms of biosafety. Several batches were produced in HEK 293T, pseudotyped for VS.V-G; the co-transfection was carried out using pLOX-Ttag-iresTK + vector VS.V-G (pMD2.G; Addgene, Watertown, MA, USA, 12259) + vector Pax2 (psPax2; Addgene, Watertown, MA, USA, 12260), using lipofectamin 3000 (Invitrogen, Madrid, Spain, L3000). The titer of the different batches was estimated by RT-qPCR using standard curves.
Primary
Bmi1+DR cells were transduced with the lentiviral vector supernatant at the indicated MOIs 1–10; cells were seeded in 6-well plates using an 80% confluency (7000 cells/cm
2). Lentiviral transduction was carried out in OPTIMEM (ThermoFisher, Waltham, MA, USA, 31985-070) supplemented with polybren (Sigma-Aldrich Inc., St. Louis, MI, USA, TR-1003) at 8 μM. Cells were maintained in the conditions previously described for
Bmi1+DR culture for 24 h, then cells were washed, and culture medium was refreshed and maintained for an additional 24 h. Finally, transduced cells were maintained in standard culture conditions, with subcultures carried out every three days until confluency was reached. The scheme of the followed procedure can be found in
Figure 2B.
For monitoring insert status by PCR, total DNA was extracted from the cells using the NucleoSpin Tissue extraction kit (Macherey-Nagel, Düren, Germany, 740952). The PCR reaction was carried out in an Applied Biosystems Veriti 96 thermoblock well (Applied Biosystems, Waltham, MA, USA) according to the following program: 10 min at 95 °C, 40 cycles of 15 s at 95 °C, 1 min at 60 °C, 30 s at 72 °C, and 7 min at 72 °C. Amplification was performed using the same primers as in the detection by RT-qPCR. For analysis, visualization of the amplificated section was performed by loading the PCR result into a 1.5% agarose gel (Condalab, Madrid, Spain, 8010.22) stained with Ethidium Bromide (Sigma-Aldrich Inc., St. Louis, MI, USA, E1610). Bands were confirmed using markers of suitable size (1 Kb DNA Plus Ladder; ThermoFisher, Waltham, MA, USA, 10787018).
4.6.2. Analysis of the Expression of Membrane Markers by Flow Cytometry
As part of the validation of the
Bmi1+DR
IMM immortalized population, we obtained an expression profile by flow cytometry for some of the markers of the membrane that characterize the
Bmi1+DR population.
Bmi1+DR
IMM cells were amplified in culture until there were 10
6 cells for each marker to analyze. The cells were trypsinized and washed twice with 1X PBS. Possible non-specific targets were blocked by 1 h incubation of cells in suspension at RT with 1X PBS + 5% BSA. After the blocking, the cell suspensions were incubated with each of the primary antibodies against the membrane markers to analyze. The marking was applied for 1 h at RT under gentle rotation to avoid precipitation of the cells. Subsequently, two washes were carried out with 1X PBS + 5% BSA and followed by labeling with the secondary antibodies under the same conditions as the primary labeling. The antibodies conjugated with fluorochromes that were used only required a marking step. The antibodies used and their concentrations of use are listed in
Table S2. Finally, fluorescently labeled cells were detected using Gallios Flow Cytometer (Beckmann Coulter, Madrid, Spain), and results were analyzed using Kaluza Analysis software version c1.2.1. (Beckmann Coulter, Madrid, Spain).
4.6.3. Evaluation of Bmi1+DRIMM Response Assays to Recombinant Proteins
With the aim of determining whether
Bmi1+DR
IMM cells keep some of the functional characteristics of primary
Bmi1+DR cells,
Bmi1+DR
IMM cells were seeded in culture plates covered with 0.1% gelatin and cultured in medium supplemented with the indicated factors and conditions described in
Supplementary Materials S2.3. Total RNA was extracted and expression levels of
Bmi1 were analyzed by RT-qPCR using the primers shown in
Table S1; modulation of
Bmi1 expression was compared with that of primary challenged
Bmi1+DR
IMM cells.
4.6.4. Dis-Immortalization: Reversal of SV40-T Immortalization by Transient Expression of Cre Recombinase
The adenoviral vector (Adeno-Cre) (SignaGen Laboratories, Frederick, MD, USA, SL100707) was prepared by the viral vector service at the CNIC (Madrid). For the vector preparation, HEK293T cells were also used, and the crude vector preparations were purified with the Adeno X-Purification Kit (Taxara, San José, CA, USA, 632249) kit, and the viral titer was established using the kit Adeno X-
TM-rapid titer Kit (Taxara, San José, CA, USA, 632250).
Bmi1+DR
IMM cells were seeded in 6-well plates to a confluency of 80% (7000 cells/cm
2). Cells were maintained for 24 h in transduction medium, and then
Bmi1+DR
IMM cells were transduced with Adeno-Cre using several MOIs (1, 2, and 5 × 10
2 infective particles for each cell); cells were incubated for an additional 72 h, in conditions equivalent to the transduction with lentiviral vector. After a step of washing the culture, medium was exchanged and the culture was kept for an additional 24 h to allow them to recover, and then the expression of the immortalizing function and selection marker (RT-qPCR and Western Blot) was monitored. The feasibility of de-immortalization was previously demonstrated [
34]. Effective Cre-dependent activity should delete the floxed cassette; in addition, treatment with ganciclovir (InvivoGen, San Diego, CA, USA, ant-nr-1). SUD-GCV) allowed us to eliminate those cells that did not delete the immortalization cassette.
Bmi1+DR
IMM cells were seeded at 7000 cells/cm
2 and, after 24 h, the negative selection with Ganciclovir (GCV, 1 μM) was added and maintained for 7 days. Associated with the expression of SV40, the cassette also expresses Timidin Kinase (TK), which metabolizes GCV, inducing cell death; this will eliminate cells that did not eliminate the immortalization cassette. The process was monitored for the reversion of the immortalization and posterior selection. Those
Bmi1+DR
IMM cells that were manipulated with the Adeno-cre and survived to GCV selection were denoted
Bmi1+DR
IMM-REV cells. The scheme of the followed procedure can be found in
Figure 3A.
4.6.5. Evaluation of Proliferative Status of Bmi1+DR Cells
Evaluation of Cell Proliferative Status through Population Doubling Rate
Population doubling rate was quantified in parallel in all treatments of immortalization for
Bmi1+DR cells compared. For this, all crops were reseeded at the same time, and in each passage, the same number of cells was seeded. The passages were performed every 3 days, and 7000 cells/cm
2 were reseeded. In the case that it was not reached, this minimum number of cells between one passage and the next, the total of the cells present in each condition was considered for the calculation of the doubling rate in the next pass. The cells were kept in culture for as long as possible, given that the control cells and those not effectively immortalized entered a stationary phase in which there was minimal or no cell proliferation. To calculate the parameter of the population doubling rate, we use the following formula:
Evaluation of Cell Proliferative Status through EdU Incorporation
To evaluate the proliferative status using the labeling EdU (5-ethynyl-2′-deoxyuridine), we incubated the cells for 12 h in standard culture medium supplemented with 10 µM EdU. Labelling for proliferative cells was performed with Click-iT EdU Alexa Fluor 647 nm Flow Cytometry Kit (Invitrogen, Madrid, Spain, C10424) following the manufacturer’s instructions. EdU+ cells were detected fluorescently within each population using the Gallios Flow cytometer (Beckmann Coulter Madrid, Spain) and results were analyzed using Kaluza Analysis software version c1.2.1. (Beckmann Coulter, Madrid, Spain).
4.6.6. Analysis of Cellular Senescence by β–Galactosidase Staining
Staining was performed against cells in a state of senescence based on the activity of β-galactosidase. The commercial Senescence β–Galactosidase kit (Cells Signaling Technology, Danvers, MA, USA, 98605) was used following the protocol indicated by the manufacturer. To determine the number of cells senescent per field, images were taken with the Olympus IX70 microscope (Olympus, Tokyo, Japan) and labeled cells were quantified Image J program version FIJI (National Institute of Health, Belthesda, MD, USA)
In addition, we evaluated several strategies to improve the survival and proliferation of
Bmi1+DR
IMM-REV dis-immortalization (see
Tables S3–S5).
4.7. Paraquat Treatments
Treatments with paraquat (N,N′-dimethyl-4,4′-bipyridinium dichloride) (Pq; Sigma-Aldrich Inc., St. Louis, MI, USA, 36541) were carried both in vivo or in vitro. For in vivo treatment,
Bmi1CreERT/+Rosa26Tomato/+ animals, 5 d post-Tx induction, were injected (i.p.) with a single dose of Pq (20 mg/kg body weight, diluted in 1X PBS), as previously described [
19]. Pq-treated animals were sacrificed, and hearts were analyzed 48 h later. For in vitro treatment, cell populations (primary or immortalized) were treated with Pq after washing once with 1X PBS and the culture medium was replaced with the corresponding culture medium without any supplement, and under culture conditions of the target cell type whose effect was analyzed, including 5 or 8 mM Pq. Treatment was maintained for 12 h and then analyzed from the different aspects. When indicated, the target population was previously labelled with Violet Tracer.
4.8. Co-Culture Experiments
To evaluate the effects of cell–cell contact between Bmi1+DRIMM cells and other cell types present in the heart through co-culture, we treated the culture surface with gelatin 0.1 or 1% and the corresponding supplements depending on the highest concentration required by the cells used in the co-culture. Subsequently, we seeded the first cell type at 25,000 cells/cm2 (HL-1, MEFs, 1 g 11 or pCECs) in its culture medium. After 8 h, we verified that these cells had adhered correctly, removed their medium culture, washed with 1X PBS, and seeded the Bmi1+DRIMM cells on top at the same density. All analyses were conducted after 12 h of co-culture in the corresponding medium and culture conditions of the target cell type whose effect was analyzed. When indicated, co-cultures were compared with non-contact cultures using transwells (Transwell Permeable Supports 0.4 μm Polycarbonate Membrane; Sigma-Aldrich Inc., St. Louis, MI, USA, 3412) to avoid cell contact.
On the one hand, we analyzed the effect of co-cultures on the survival of the different cell types studied against exposure to severe oxidative damage (Pq treatment,
Section 4.7). Because the low size of the fluorescence marker YFP in
Bmi1+DR
IMM cells was significantly diffused after Pq treatment, confusing results (see
Figure S4), we used cells previously labelled with the CellTrace™ Violet reagent (ThermoFisher, Waltham, MA, USA, C34557) at a concentration of 5 μM in a ratio of 1 million cells/mL 1X PBS, incubating for 20 min in darkness at 37 °C. After the administration of Pq, co-culture was maintained with the co-culture for an additional 12 h, before analysis. Dead cells were analyzed for propidium iodide staining (PI; Abcam) or DAPI staining. After the Pq treatment, dead cells labeled with propidium iodide (PI; Abcam) or DAPI (Beckman Coulter) within each population (Violet-/Violet+) were detected using the Gallios Flow Cytometer (Beckmann Coulter) and results were analyzed using Kaluza Analysis software version c1.2.1. (Beckmann Coulter, Madrid, Spain).
On the other hand, we analyzed gene expression in co-cultures of Bmi1+DRIMM cells with pCECs. First, we separated the different cell types present in each co-culture. To accomplish this, we trypsinized the cells and separated them by FACS. This technique allowed the separation of fluorescently labeled Bmi1+DRIMM (YFP+) cells from the negative without any type of labeling. Once the different fractions were separated, we extracted total RNA and analyzed the variations in expression with respect to the Bmi1+DRIMM mono-culture as control by RT-qPCR.
4.9. Autophagy Evaluation by LC3B Detection, Difference between Total and Canonical Autophagy
Direct contact co-culture of pCECs with
Bmi1+DR cells was performed as described (see
Section 4.10). LC3B detection was carried out using the Guava Autophagy LC3-antibody-based assay Kit (Luminex, Austin, TX, USA, FCCH100171), following the instructions described by the manufacturer. Due to the detection of LC3B antibody (488 nm),
Bmi1+DR
IMM cells (YFP+) were not suitable; in this case, we performed co-culture of pCECs with primary
Bmi1+DR cells (labelled prior with Violet tracer). Co-culture and mono-culture, as control, were maintained for 12 h under
Bmi1+DR culture conditions using
Bmi1+DR culture medium supplemented with bafilomycin (10 μM), chloroquine (40 μM), and without supplement as control. Treatments with bafilomycin and chloroquine, as previously described [
78], allowed us to discriminate between canonical and non-canonical autophagy. Anti-LC3B 488 nm detection (% Max intensity) was performed using Gallios Flow Cytometer (Beckmann Coulter, Madrid, Spain) and results were analyzed using Kaluza Analysis software version c1.2.1. (Beckmann Coulter, Madrid, Spain). Total autophagy was calculated as the ratio of Anti-LC3B detection of [(Chloroquine treated cells−Untreated cells)/Untreated cells], Canonical autophagy as the ration of Anti-LC3B detection of [(Bafilomycin treated cells−Untreated cells)/Untreated cells], and finally, non-canonical autophagy as the difference between total and canonical autophagy.
4.10. Metabolism Activity by Seahorse Analysis
Direct contact co-culture of pCECs with
Bmi1+DR
IMM cells was performed as described (see
Section 4.8). Co-culture was maintained for 12 h, then pCECs were separated from the co-culture by magnetic separation using the MACS Neonatal Cardiac Endothelial Cell Isolation Kit.
Bmi1+DR
IMM cells were seeded in specific cell culture microplates and metabolic activity was carried out by Agilent Seahorse XF96 kit (Agilent technologies, Madrid, Spain, V3-PS TC-Treated, 101085-004). Results were analyzed in XF96 Analyzer obtaining the percentage of oxygen consumption rate (OCR) and the percentage of extracellular acidification rate (ECAR), as previously described [
79].
4.11. Statistical Analysis
Statistical analyses were carried out with GraphPad Prism 7.0 software. For the study of data composed of a number of experimental samples greater than 15 (n > 15), the distribution was analyzed using the Shapiro–Wilk test, considering a normal distribution when p ≥ 0.05. In the analysis of experiments composed of two conditions, the Mann–Whitney U-Test was used. For experiments in which multiple conditions were analyzed, the Kruskal–Wallis test followed by Dunn’s post-test was used in the case of samples with parametric non-distribution or the one-way ANOVA test followed by Bonferroni post-test in the case of comparisons with parametric distribution. Significant differences were considered in the experiments that had a p-value less than 0.05 (* p < 0.05, ** p < 0.01, *** p < 0.001).