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. 2025 Jan 23;20(2):102394. doi: 10.1016/j.stemcr.2024.102394

Microvessel co-transplantation improves poor remuscularization by hiPSC-cardiomyocytes in a complex disease model of myocardial infarction and type 2 diabetes

Xuetao Sun 1, Jun Wu 1, Omar Mourad 1,2, Renke Li 1,3, Sara S Nunes 1,2,3,4,5,6,∗
PMCID: PMC11864147  PMID: 39855203

Summary

People with type 2 diabetes (T2D) are at a higher risk for myocardial infarction (MI) than age-matched healthy individuals. Here, we studied cell-based cardiac regeneration post MI in T2D rats modeling the co-morbid conditions in patients with MI. We recapitulated the T2D hallmarks and clinical aspects of diabetic cardiomyopathy using high-fat diet and streptozotocin in athymic rats, which were then subjected to MI and intramyocardial implantation of human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) with or without rat adipose-derived microvessels (MVs). hiPSC-CM alone engrafted poorly. Co-delivery of hiPSC-CMs with MVs yielded a smaller infarct area and a thicker left ventricle wall. Additionally, MVs robustly integrated into the infarcted hearts, improved the survival of hiPSC-CMs, and improved cardiac function. MV-conditioned media also promoted hiPSC-CM maturation in vitro, increasing cardiomyocyte (CM) size in an interleukin (IL)-6-dependent manner. Given the availability of MVs from human adipose tissue, MVs present great translational potential for the treatment of heart failure in people with T2D.

Keywords: T2D, diabetic cardiomyopathy, microvessel, myocardial infarction, hiPSC-cardiomyocytes, cell transplantation, cardiac regeneration

Graphical abstract

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Highlights

  • •

    HFD feeding and STZ injection in RNU nude rats reproduce several hallmarks of T2D

  • •

    MVs promote the survival and hypertrophy of hiPSC-CMs in a complex disease model

  • •

    IL-6 depletion from the MV-conditioned media prevents PSC-CM hypertrophy in vitro

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    Co-transplantation of MV and hiPSC-CM improves heart function post MI in T2D rats

In this article, Nunes Vasconcelos and colleagues show that co-delivery of adipose-derived microvessels and hiPSC-cardiomyocytes improves cardiomyocyte survival, graft size, and cardiac function in a complex disease model that better reflects the comorbid conditions in the clinic (T2D + MI). hiPSC-cardiomyocytes undergo hypertrophy when treated with microvessel-conditioned media in vitro, which is abolished when IL-6 is depleted from media.

Introduction

A promising treatment for myocardial infarction (MI) lies in cell-based regenerative therapies in which human pluripotent stem cell-derived cardiomyocytes (hPSC-CMs) are transplanted into the infarcted area to replace the cardiomyocytes (CMs) lost post MI (Chong et al., 2014; Laflamme et al., 2007; Shiba et al., 2012; Sun et al., 2020). While hPSC-CM transplantation has been tested in animal models of MI, most studies are performed in healthy animals that lack the comorbid conditions that contribute to the development of MI and may impede endogenous and cell-based therapies. The use of animal models without co-morbidities has contributed to a failure to translate findings into the clinic and argues for the use of more relevant disease models in pre-clinical studies.

The incidence of MI in people with diabetes mellitus (DM) is significantly higher than in the age-matched non-diabetic population. DM is a major metabolic disorder, and type 2 diabetes (T2D) is the predominant form, accounting for 90% of cases worldwide (Chatterjee et al., 2017). People with diabetes are at an increased risk for cardiovascular diseases, which are the leading cause of morbidity and mortality in this cohort, accounting for an estimated 80% of all deaths in people with diabetes in North America (Glass et al., 2010), including those from MI and heart failure. DM is also known to impair vascularization, which exacerbates the already extensive hypoxia-driven hPSC-CM death reported in MI models in healthy animals. Therefore, the development of strategies to restore cardiac function of the infarcted diabetic heart may present great therapeutic potential.

We have recently established a preclinical vascularization strategy to support the survival and engraftment of transplanted human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) in rat hearts post MI using primary microvessels (MVs) isolated from the rat adipose tissue (Sun et al., 2020). These MVs retain endothelialized lumens and perivascular cell coverage and can rapidly inosculate with the hosts’ vasculature to form perfused vascular networks after transplantation (Aghazadeh et al., 2021; Altalhi et al., 2017; Sun et al., 2020). Co-transplantation of MVs with hiPSC-CMs accelerated blood perfusion of hiPSC-CM grafts post MI in nude athymic rats (RNU), enhanced the survival of the transplanted CMs by 6-fold, and led to cardiac functional recovery (Sun et al., 2020).

To increase the clinical relevance of our approach, we tested the MV-based vascularization in an MI-model in T2D rats. First, we created a T2D model in animals that are immunodeficient to allow xenograft studies (Festing et al., 1978) by feeding RNU rats a high-fat diet (HFD) followed by injection of a single dose of streptozotocin (STZ) (Reed et al., 2000). Animals showed the hallmarks of T2D, including the cardinal features of diabetic cardiomyopathy.

We then tested the efficiency of implantation of hiPSC-CM only or with MVs in T2D rats post MI. We report that hiPSC-CMs show poor survival in the infarcted hearts of T2D rats and that MVs robustly integrated into the infarcted coronary vasculature, enhanced the survival of hiPSC-CMs, and promoted cardiac functional recovery.

Results

Induction and characterization of T2D in RNU rats

To test the efficacy of the MV-based therapeutical strategy in a more clinically relevant model of MI in diabetic animals, we first developed and characterized a T2D model in RNU rats that are suitable for xenograft cell transplantation using HFD and STZ (HFD-STZ) (Figure S1A). RNU rats were randomized into a control group (normal chow) or an HFD (45% fat, 20% protein, and 35% carbohydrate)-STZ group. STZ (35 mg/kg i.v.) or vehicle was administered 8 weeks after HFD feeding. Assessments were done longitudinally and at week 9 (endpoint). There were no differences in body weight between control and HFD-STZ cohorts over time (Figure S1B). At week 8, the HFD-STZ group showed a mild but significant increase in blood glucose levels pre-STZ administration (7.7 ± 0.3 mM vs. 5.8 ± 0.2 mM in control) (Figure S1C).

To test if HFD feeding induced insulin resistance, we performed an insulin tolerance test at the 8-week time point prior to STZ injection. Fasting plasma insulin levels were significantly higher in HFD-STZ rats compared to normal chow control (Figure S1D). The degree of insulin resistance in the HFD-STZ group was significantly higher than in control, as shown by the assessment of homeostatic model assessment of insulin resistance (Figure S1E). Additionally, insulin sensitivity was reduced in HFD-STZ animals compared to control rats, as shown by the blood glucose curves (Figures S1F and S1G).

Diabetes was exacerbated at week 9, after the administration of STZ when blood glucose levels increased more substantially (23.6 ± 1.4 mM vs. 5.5 ± 0.3 mM in control) (Figure S1C).

Assessment of blood lipids at week 8 revealed significantly increased high-density lipoprotein (HDL)-cholesterol, total cholesterol, and triglycerides in the HFD cohort relative to control (Figures S1H, S1J, and S1K). No differences were observed for low-density lipoprotein-cholesterol between groups (Figure S1I).

The aforementioned changes were in line with T2D models in immunocompetent rats (Reed et al., 2000; Srinivasan et al., 2005; Zhang et al., 2008) and showed that we successfully established a model of T2D in RNU rats. Next, we investigated the presence of potential cardiovascular alterations as a consequence of T2D.

Functional analysis via echocardiography revealed that prior to STZ injection (week 8), fractional shortening (FS) was already significantly lower in HFD animals (42.9% ± 1.6%) compared to control (48.4% ± 1.3%) (p < 0.001, Figure S2A). FS remained significantly lower in HFD-STZ rats (44.1% ± 2.1%) compared to healthy controls (47.8% ± 1.4%) after STZ injection (p = 0.002, Figure S2A). In addition, despite an initial significant increase in left ventricular internal dimension in diastole (LVIDd) and left ventricular internal dimension in systole (LVIDs) in HFD rats at week 8, prior to STZ injection (Figures S2B and S2C, week 8), after treatment with STZ, there were no differences in LVIDd (8.1 ± 0.3 mm) compared to control (7.7 ± 0.5 mm) (p = 0.110, Figure S2B). LVIDs was still significantly higher in HFD-STZ rats (4.5 ± 0.3 mm) compared to normal chow control (4.0 ± 0.3 mm) (p = 0.010, Figure S2C). These results showed mild cardiac dysfunction in diabetic animals that was a consequence of HFD and the presence of insulin resistance in RNU rats, prior to fully established diabetes.

Consistent with diabetes-induced cardiac pathology, we observed cardiac hypertrophy in HFD-STZ rats at week 9 as demonstrated by (1) the significant increase in the ratio of heart weight by tibial length (p = 0.005, Figure S2D), (2) increased left ventricular wall thickness (p = 0.005, Figures S2E and S2F), and (3) increased cardiomyocyte size as assessed by the surface area of wheat germ agglutinin (WGA)+ cardiac troponin T (cTnT)-stained sections (Sun et al., 2020) in HFD-STZ rats compared to age-matched controls (p = 0.001, Figures S2G and S2H).

We also investigated the presence of cardiac fibrosis and microvascular rarefaction as these have been associated with diabetic cardiomyopathy (Hinkel et al., 2017; Jia et al., 2018). Picrosirius red staining revealed a significant increase in collagen deposition in HFD-STZ rats compared to controls at week 9 (p = 0.05, Figures S2I and S2J), indicative of fibrosis. Vessel density was slightly reduced (18%) in the HFD-STZ rats (1,743 ± 67/mm2) compared to controls (2,062 ± 180/mm2), but not at a significant level (p = 0.149, Figures S2K and S2L).

In summary, in line with the HFD-STZ model in immunocompetent rats, the HFD-STZ treatment of RNU rats recapitulates key features of T2D. These will be referred to as T2D rats henceforth.

MVs enable the survival of hiPSC-CMs and heart remuscularization in a complex disease model of MI in T2D rats

Next, we investigated whether co-transplantation of MVs and hiPSC-CMs would lead to an increase in hiPSC-CM survival and improve cardiac remuscularization post MI in T2D rats (Figure 1). MI was induced by left coronary artery (LCA) ligation, as previously described (Sun et al., 2020). Two weeks post MI, animals with FS < 20% or >35% were excluded from the study and rats were randomized into three groups: MI-only (no treatment), CM-only (hiPSC-CMs), and CM + V (hiPSC-CMs and MVs). We delivered 10 × 106 hiPSC-CMs (>95% purity) only or with 5,000 MVs (∼2 × 105 cells, <2% of total cells delivered) in 100 μL of collagen type I gel via direct transepicardial injection (Figure 1), as in the study by Sun et al. (2020). Heart function, cardiac structure, and cardiomyocyte engraftment were assessed 4 weeks post cell transplantation (Figure 1).

Figure 1.

Figure 1

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Schematic of the experimental protocol

RNU rats were fed a high-fat diet (HFD) for 8 weeks before receiving a single dose (35 mg/kg body weight) of STZ (i.v.). Diabetic rats underwent LCA ligation (MI, week 0) followed by intramyocardial injection of 10 × 106 hiPSC-CMs (CM-only) or with 5 × 103 MVs harvested from adipose tissue (CM + V) 2 weeks after MI. Cardiac function was assessed longitudinally via Echo and at endpoint by PV loop analysis. Histology was performed 4 weeks (W4) after cell delivery. BG, blood glucose; Echo, echocardiography; LCA, left coronary artery; LV, left ventricle; PV loop, pressure-volume loop; STZ, streptozotocin.

At 4 weeks after transplantation, the infarct area was significantly smaller (p < 0.05) in CM + V, but not in the CM-only group, compared to the MI-only control (Figures 2A–2C). Left ventricle wall thickness was larger in the CM + V group compared to CM-only and MI-only groups (Figures 2B and 2D). The infarct border zone in the CM + V was significantly thicker compared to CM-only or MI-only groups (Figure S3A). Collagen in the border zone was significantly lower in the CM + V group compared to CM-only or MI-only groups (Figure S3B). CM + V rats had significantly lower heart weight by tibial length (Yin et al., 1982) compared to CM-only or MI-only rats (p < 0.05, Figure 2E), suggesting a slowed progression in cardiac hypertrophy.

Figure 2.

Figure 2

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MVs improve infarcted heart remuscularization in diabetic rats post MI

(A) Representative images of rat hearts 4 weeks after implantation showing MI-only (control), CM-only, and CM + V. Scale bars, 10 mm.

(B) Representative hematoxylin and eosin (H&E) (left) and picrosirius red (right) staining of short-axis sections. Scale bars, 2 mm.

(C) Assessment of infarct area in MI-only, CM-only, and CM + V groups. MI-only (n = 4), CM-only (n = 8), and CM + V (n = 11) (n, number of rats). Data are presented as mean ± SEM.

(D) Comparison of left ventricle wall thickness in MI-only, CM-only, and CM + V groups. MI-only (n = 4), CM-only (n = 8), and CM + V (n = 11) (n, number of rats). Data are presented as mean ± SEM.

(E) Assessment of heart weight and tibia length ratios in MI-only, CM-only, and CM + V groups. MI-only (n = 3), CM-only (n = 4), and CM + V (n = 5) (n, number of rats). Data are presented as mean ± SEM. One-way ANOVA with Bonferroni multiple comparisons test was used for multiple groups in (C)–(E).

(F) Representative images of heart sections containing grafts at week 4 stained with cTnT and a human-specific anti-Ku80 antibody with nuclei counterstained by Hoechst in CM-only and CM + V groups. Scale bars, 50 μm.

(G) Quantification of total Ku80+ cells/heart section of CM-only and CM + V groups 4 weeks after implantation, indicative of hiPSC-CM survival. CM-only (n = 8) and CM + V (n = 11) (n, number of rats). Shapiro-Wilk test followed by Mann-Whitney rank-sum test. Data are presented as mean ± SEM.

hiPSC-CMs grafts were identified using a human-specific, anti-Ku80 antibody (Allard et al., 2014; Sun et al., 2020) and cTnT (Figure 2F) or myosin heavy chain (MHC) (Figure S4A). Consistent with previous reports, the success rate of hiPSC-CM engraftment into ischemic heart was low, with only 13% of the hearts having detectable grafts. In contrast, the co-delivery of hiPSC-CMs and MVs increased the engraftment detection rate to 73%. Co-implantation with MVs led to a 118-fold increase in hiPSC-CM number (Ku80+/cTnT+ cells) (Figures 2F and 2G) and a 231-fold increase in graft size (Figure S5B). The graft area normalized to the left ventricular (LV) area (graft size as percentage of LV area) was significantly higher in animals receiving CM + V compared to CM-only (Figures S4C and S4D). In addition, in line with the reported cardiac hypertrophy, the total LV area was larger in diabetic rats (Figure S4E).

Increase in CM graft size by MVs is not due to high glucose

When comparing the size of the grafts at 4 weeks post transplantation between the T2D rats and the previous study where MI was performed in non-diabetic rats, we found that the size of CM + V grafts was larger in T2D + MI rats than those in non-diabetic MI ones (Sun et al., 2020). Specifically, there was a 5.3-fold increase in graft size by MV co-transplantation vs. CM-only in non-diabetic rats (Sun et al., 2020) and a 231-fold increase in T2D rats (Figure S4B). This was unexpected as it is in general more difficult to form new blood vessels in diabetes as high glucose negatively impacts endothelial cells. Thus, we sought to determine if the larger graft sizes in diabetic rats were due to high glucose as hiPSC-CMs rely on glucose as the main source of energy. Therefore, we implanted CM + V in collagen gel subcutaneously into diabetic or non-diabetic mice. There were more cardiomyocytes (cTnT+), and the cTnT+ area was significantly larger in grafts implanted into non-diabetic mice than in diabetic ones (p = 0.018, Figures S4F–S4H), suggesting that the increase in graft size seen in the diabetic hearts post MI is likely not due to the high-glucose environment.

Co-transplantation with MVs promote heart function recovery

Similar to MI-only, the transplantation of hiPSC-CM-only did not improve FS (Figure 3A) compared to baseline. However, co-transplantation of CM + V reversed the FS decline, improving function when compared to CM-only (Figures 3A and 3B). There were no differences in LVIDd in CM + V compared to MI-only or CM-only (Figures 3C and 3D). LVIDs was significantly better in CM + V compared to MI-only, but not to CM-only (Figures 3E and 3F), suggesting that the functional improvement derived mainly from the systolic change. These data demonstrate the superiority of MVs in supporting cardiomyocyte transplantation and in improving heart function.

Figure 3.

Figure 3

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Co-transplantation of MVs and hiPSC-CMs restores ventricular function in diabetic rat post MI

(A–F) Longitudinal assessment of heart function by echocardiography (A–F). MI-only (n = 6), CM-only (n = 8) and CM + V (n = 11) (n, number of rats). Each dot represents one animal. Fractional shortening values (A), LVIDd (C), and LVIDs (E) for the pre-injection baseline and 4 weeks after injection. Differences in fractional shortening (B), LVIDd (D), and LVIDs (F) between baseline and W4 after injection. NS, not significant. Two-tailed paired t test was used for comparison of cardiac function within groups between pre-injection baseline and W4 time point. One-way ANOVA with Bonferroni multiple comparisons test was used for multiple groups.

(G) Representative PV loop 4 weeks after transplantation.

(H) Ejection fraction 4 weeks after transplantation. MI-only (n = 6), CM-only (n = 8), and CM + V (n = 11) (n, number of rats). ∗, p < 0.05, ∗∗p < 0.001 vs. MI-only, ##, p < 0.001 CM-only vs. CM + V. One-way ANOVA with Bonferroni multiple comparisons test were used for multiple groups. Data are presented as mean ± SEM.

Pressure-volume (PV) loops at 4 weeks after treatment showed a higher end-systolic pressure (ESP) and smaller systolic volume in CM-only hearts compared to MI-only controls, suggesting an improved cardiac function (Figure 3G). Co-implantation of MVs further improved ESP and systolic volume (Figure 3H). Consistent with echocardiographic data, PV analysis showed a significant improvement in ejection fraction (EF) in CM + V group (∼36%) compared to both CM-only (∼26%, p < 0.001) and MI-only (∼21%, p < 0.001) groups (Figure 3H). Similarly, Tau (the isovolumetric relaxation time constant), maximal slope of systolic pressure increment (dP/dt max), diastolic decrement (dP/dt min), end-systolic volume (ESV), and ESP, except end-diastolic volume (EDV), were all further significantly improved in the CM + V group compared to the CM-only group (Figure S5).

MVs increase vessel density, integrating long term into the cardiac vasculature

Next, we investigated whether the co-transplantation of hiPSC-CMs with MVs led to an increase in graft vascularity by quantifying CD31+ vessels 4 weeks after treatment (Figure 4A). We found a 13.7-fold increase in CD31+ vessel area in CM + V grafts compared to the CM-only group (Figure 4B). Vessel area in CM + V grafts was 1.9-fold lower than in the uninjured native heart tissue (Figure 4B). The vessels in the CM + V graft displayed blood perfusion as shown by the presence of red blood cells in their lumens (Figure 4C). There were no obvious signs of vessel degeneration. To determine the origin of the vessels in the CM + V grafts, we used MVs isolated from rats that ubiquitously express GFP. We detected a high percentage of GFP+ vessels in the graft (Figure 4D), indicating donor vessel persistence. Vessel density assessed by intrinsic GFP expression (Figure 4D) or by staining with CD31 (total vessels) was similar (Figure 4E), indicating that the majority of vessels in CM + V grafts are of donor origin. In addition, vessels of donor origin (GFP+) were perfused as shown by the quantification of lectin perfused intravenously into the host rat (Figures 4D and 4F).

Figure 4.

Figure 4

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MVs improve vessel density, persist long term, and integrate into the cardiac vasculature forming stable grafts

(A) Representative heart sections of CM only and CM + V grafts 4 weeks after delivery stained with anti-CD31 identifying total (host and donor) endothelial cells. Scale bars, 50 μm.

(B) CD31 area assessed in CM only and CM + V grafts and native heart tissue (remote area). CM only (n = 1), CM + V (n = 4), and native tissue (n = 5) (n, number of rats). ∗∗p < 0 001 vs. CM only; ##p < 0 001, native tissue vs. CM + V. Data are presented as mean ± SEM.

(C) Higher magnification images of grafts 4 weeks after delivery stained with anti-CD31 antibody and counterstained with H&E showing presence of red blood cells (indicative of blood perfusion).

(D–F) Vessel persistence was assessed week 4 after implantation of vessels harvested from rats that ubiquitously express GFP.

(D) Representative images of two consecutive heart sections showing hiPSC-CMs (green, left) and donor vessels (green, right), and these vessels are perfused by host circulation (right). Scale bar, 50 μm.

(E) Donor vessel density (GFP+ vessels) (n = 5) and total (CD31+) (n = 4) vessel density in week 4 grafts. n, number of rats. Data are presented as mean ± SEM.

(F) Percentage of perfused donor vessel in week 4 grafts. n = 5 rats. Data are presented as mean ± SEM.

MVs induce hiPSC-CM hypertrophy

Analysis of hiPSC-CM cell surface area 4 weeks after cell delivery showed that hiPSC-CMs in the CM + V rats were significantly larger compared to cardiomyocytes from CM-only rats, suggesting an MV-specific induction in hiPSC-CM hypertrophy (p = 0.001, Figure 5). To test whether MV-secreted factors promote CM hypertrophy, we treated human embryonic stem cell-derived cardiomycetes (hESC-CM) in vitro with different concentrations of the conditioned media of MVs. Consistent with in vivo findings, the conditioned media of MVs significantly increased the cell surface area of hESC-CMs, in a dose-dependent manner (Figures 6A and 6B).

Figure 5.

Figure 5

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MVs promote hiPSC-CM hypertrophy in vivo

(A) Representative images of CM-only and CM + V grafts 4 weeks after treatment stained with WGA and anti-human Ku80. Scale bars, 50 μm.

(B) Violin plots showing quantification of hiPSC-CM surface area. CM only (n = 1); CM + V (n = 4) (n, number of rats).

Figure 6.

Figure 6

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MVs secreted hypertrophic factors promoting hESC-CM hypertrophy in vitro

(A) hESC-CMs were incubated with the indicated concentrations of conditioned media for 1 week prior to immunofluorescence (cTnT and Hoechst). Scale bar, 100 μm.

(B) Quantification of the cell surface area. n = 3 independent experiments. Data are presented as mean ± SEM.

(C) Detection of hypertrophic factors secreted by MVs in culture in media with different concentrations of glucose using cytokine array.

(D) Antibody-mediated depletion of IL-6. n = 3 independent experiments. Data are presented as mean ± SEM.

(E) hESC-CM cultured in MV media before and after IL-6 depletion. Cells were stained with α-actinin with nuclear counterstained by Hoechst. Scale bar, 100 μm.

(F) Quantification of cell surface area of the hESC-CM. n = 3 independent experiments. ∗, p < 0.05, ∗∗p < 0.001, ns, not significant. One-way ANOVA with Bonferroni multiple comparisons test were used for multiple groups. Data are presented as mean ± SEM.

The average sarcomere length in the CM + V grafts was 1.79 ± 0.03 μm (Figures S6A and S6B). In vitro, there was no difference in sarcomere length in hiPSC-CMs treated with MV-conditioned media (Figure S6C). Assessment of CM proliferation showed that 5.9% of human CMs in CM + V grafts were positive for Ki67 (Figures S6D and S6E).

MV-dependent hESC-CM hypertrophy is mediated by IL-6 in vitro

Analysis of the conditioned media of MVs in culture in normal (5 mM) and high (15 or 25 mM) glucose demonstrated that high glucose did not disrupt the secretion of hypertrophic cytokines by MVs in culture (Figure 6C). Further, antibody depletion of interleukin (IL)-6 from the MV-conditioned media was sufficient to prevent CM hypertrophy (Figures 6D–6F).

Discussion

To study the compounded effects of T2D myocardial dysfunction and MI in cardiac regeneration, we first generated an HFD-STZ-induced T2D model using RNU rats that are amenable for xenograft studies. We showed that RNU rats display several hallmarks of T2D, including cardiac dysfunction. Though there are several versions of the HFD-STZ to induce diabetes in rats, varying in length of HFD feeding and in STZ dose and number of injections, the protocols are similar: HFD feeding for short (2–4 weeks) or long (over 3 months) periods followed by STZ (Hu et al., 2013; Reed et al., 2000; Srinivasan et al., 2005; Zhang et al., 2008). Given practical and financial considerations, our chosen regimen was intended to be short while successfully showing key hallmarks of the disease.

There are great variations in the STZ protocols. Though there is no consensus on the STZ treatment in modeling T2D, we elected to administer a low dose of STZ to decrease the potential of toxicity (Szkudelski, 2001), which was effective in inducing hyperglycemia. It should be noted that STZ treatment led the RNU rats to transition from an insulin-resistant state showing mildly elevated blood glucose levels to blood glucose levels of more established T2D within 1 week, which might not precisely mimic the progression of the disease in humans and is therefore a limitation of the model.

It is well known that diabetes affects the heart and can lead to diabetic cardiomyopathy. We showed that HFD-STZ treatment induced myocardial thickening, cardiomyocyte hypertrophy, and increased collagen deposition (i.e., fibrosis) compared to healthy controls by week 9. Assessment of myocardial microvascular density revealed a small but not significant decrease in vessel density in diabetic rats. However, it is likely that vascular rarefaction would take longer to appear as seen in chronic T2D. Overall, our findings are in line with the hallmarks of established diabetic cardiomyopathy. Our data also showed elevated total cholesterol and triglycerides. These changes are concurrent with the structural and functional cardiac changes. However, HDL-cholesterol, associated with a lower risk of heart disease, was also significantly elevated. Similar results have been reported elsewhere in a different strain (Loai et al., 2021).

The immune system is a key element in T2D and contributes to its progression from pre-diabetes to fully established T2D in different ways (Zhou et al., 2018). Although RNU rats are immunocompromised, it is not surprising that they still showed several hallmarks of T2D as RNU rats, although lacking a normal thymus and functionally mature T cells, have normal numbers and the full range of immune cells, including B and natural killer cells, monocytes, and macrophages (Festing et al., 1978; Rolstad, 2001).

However, this model has several limitations. First, sex is a component of rigor and reproducibility in animal studies, and female animals should be included in the future to provide a more efficient and effective experimental design. Second, we used 6–7 weeks as the initial age of the rats, similar to other HFD-STZ models (Reed et al., 2000; Sahin et al., 2007; Si et al., 2012). Though the prevalence of T2D in children and youth is increasing, the disease is still much more prevalent in older age groups (Saeedi et al., 2019). However, starting with younger animals allowed us to focus the MI studies on adult animals and not have to take into consideration potential additional effects of aging.

Once the model was characterized, we investigated whether the delivery of hiPSC-CM would lead to engraftment and if co-transplantation of MVs would improve hiPSC-CM survival and enhance cardiac functional recovery post MI. We demonstrated that hiPSC-CM survival was poor in the infarcted hearts of T2D rats, at a similar extent as in non-diabetic rats (Sun et al., 2020). Co-delivery of MVs with hiPSC-CMs significantly improved the survival of hiPSC-CMs and increased graft size compared to CM-only.

Interestingly, comparison of CM + V graft sizes in this study with our previous studies in non-diabetic rats showed that the total number of hiPSC-CMs/section was larger in T2D than in non-diabetic rats. This goes against what we anticipated, given that this model would be more severe. To better compare these separate studies, we assessed graft size as a function of LV area. This showed that CM + V grafts occupied ∼1% of the LV area in both non-diabetic and T2D cohorts, suggesting graft of similar sizes between studies. This would also be unexpected and shows that MVs also provide support for transplanted cells in complex diseases. Of note, LVs were larger in T2D rats due to hypertrophy, suggesting that size of the grafts (%) in the T2D cohort might be underestimated. Given these unexpected results and that hiPSC-CMs are highly glycolytic compared to mature CMs that rely preferentially on fatty acid oxidation, we tested if transplantation in high-glucose environment would provide a survival benefit to hiPSC-CMs. We showed that the transplantation of CM + V subcutaneously into diabetic or non-diabetic mice led to significantly larger grafts in healthy mice than in diabetic ones. This suggests that it is unlikely that the high-glucose environment offered a potential metabolic advantage to the survival of the immature hiPSC-CM.

Another limitation of this study is that the permanent coronary occlusion MI model does not reflect the MI reperfusion in patients, though it produces robust remodeling response and large effect size. As such, the low engraftment rate of the CM-only group and the enhanced effects of CM + V might be exaggerated.

It is well established that diabetes/high glucose causes defects in vascularization, potentially making cell transplantation more challenging. We and others (Aghazadeh et al., 2021; Nalbach et al., 2021) have shown that MV-based vascularization strategies are applicable in diabetic conditions with evidence showing high persistence of donor MVs. This study supports high vessel persistence in a more complex disease model of T2D + MI, which suggests the presence of more mature and stable vessels. However, the long-term effects (beyond 4 weeks) of exposure to high glucose in the integrity and fate of transplanted vessels remain to be determined.

In terms of potential for translation, an autologous approach for MV transplantation would be desired and would avoid immunosuppressants. Though we show here that MVs improve cell survival and help cardiac functional recovery post MI and that MVs from T1D animals still show regenerative potential, albeit with a delay in perfusion (Altalhi et al., 2019), there are concerns related to the loss of regenerative potential of MVs harvested from diabetic donors. This would prevent the use of an autologous approach for transplantation into people with diabetes. This could limit clinical application to patients who do not have microvascular diseases (e.g., patients with other co-morbid conditions such as atherosclerosis that affect predominantly large vessels). Thus, more studies are needed to evaluate the potential therapeutic use of MVs from diabetic donors.

We used MVs obtained from the rat epididymal fat pad, which have been shown to promote cell survival and functional recovery of hPSC-CM in non-diabetic rats (Sun et al., 2020) and T2D rats (this study). However, this fat source may not be easily translatable to human and would require more accessible and larger adipose tissue sources. We have previously successfully used MVs harvested from the human subcutaneous fat layer via liposuction (Strobel et al., 2021) for the transplantation of human pancreatic progenitors and human islets in mouse models of T1D (Aghazadeh et al., 2021). Given the complexity of diabetes, stem cell-based therapies may be combined with other strategies to treat diabetes, including antidiabetic drugs (Bruin et al., 2015) or other sustained methods (Beckerman et al., 2021).

We examined sarcomere length, an indicator of cardiomyocyte maturation. The average sarcomere length of 1.79 μm found in the CM + V group is suggestive of immature cardiomyocytes (Dhahri et al., 2022). Unfortunately, since only 13% of the rats that received CM-only had grafts and the grafts were very small, there were not enough samples for assessing sarcomere length in the CM-only cohort, preventing the direct comparison between groups. In vitro, there was no difference in sarcomere length between hPSC-CMs treated with MV-conditioned media or control media. hPSC-CM proliferation in CM + V grafts was similar between non-diabetic (Sun et al., 2020) and T2D cohorts. Taken together, the results suggest that while there might be some maturation, shown by the increased CM size, maturation was not observed across all parameters. This is in line with previous reports that structural maturation can happen in response to mechanical stimulus but metabolism remains immature (Gomez-Garcia et al., 2021; Protze et al., 2019). This is not unexpected given that the diabetic environment is known to negatively affect CM health.

We have also demonstrated that the in vivo maturation of hPSC-CMs mediated by MVs can be replicated in vitro using the conditioned media from MVs, confirming the paracrine nature of this phenomenon. We have also implicated IL-6 as a potential mediator. IL-6 is a pleiotropic cytokine expressed by multiple cell types including immune cells, vascular endothelial cells, smooth muscle cells, fibroblasts, and cardiomyocytes (Plenz et al., 2002; Schmidt-Arras and Rose-John, 2016). IL-6 has been shown to induce cardiomyocyte hypertrophy (Hirota et al., 1995). A widespread increase in IL-6 levels is an indicator of worsening cardiac function and is associated with a poor prognosis in patients (Hirota et al., 2004; Torre-Amione et al., 1995). Here, we found no evidence of adverse functional effects of MV co-transplantation suggesting that IL-6 secretion by MVs may be local and have no widespread deleterious effects. It should be noted that the transplanted MVs persist at least for 4 weeks, and potentially longer as we have shown in T1D (Aghazadeh et al., 2021). It is unknown whether grafted MVs serve as a source of IL-6 long-term, in a constant manner or if only a brief amount of time. This also applies to other angiocrine factors secreted by MVs (Sun et al., 2020). In the future, it would be interesting to determine their individual and/or collective secretory potential over time—which could be assessed by single-cell transcriptomics.

Though people with diabetes are potential candidates to receive hPSC-CM transplantation to improve cardiac function, it is possible that they may not respond as effectively due to the altered biology of the diabetic hearts (Tan et al., 2020). Our study showed that while the success of hPSC-CM transplantation is low, MVs were effective in promoting hPSC-CM survival and functional improvement in T2D rats post MI, indicating that the MV therapy is robust and amenable for regenerative medicine application in complex diseases.

Experimental procedures

Animal

Male athymic rnu/rnu rats (6–7 weeks, Charles River) weighing 160–200 g were used. All the animal treatments and the experimental procedures were approved by the Animal Care Committee of University Health Network (Toronto, Ontario, Canada) under AUP4153 and conducted in accordance with the Guide for the Care and Use of Laboratory Animals published by the National Institutes of Health and Canadian Council on Animal Care guidelines. All rats were housed in plastic cages in standard laboratory conditions (20°C, 56% RH). Rats were maintained under a 12-h light/dark cycle (6 a.m./6 p.m.) and had ad libitum access to food and water.

Induction of T2D by HFD feeding and STZ at low dose

The normal chow consisted of 3.1 kcal/g, comprising 17% calories from fat, 25% from protein, and 58% from carbohydrate (LM-485, Teklad, Envigo, Madison, WI, USA). The HFD consisted of 4.7 kcal/g, comprising 45% calories from fat, 20% from protein, and 35% from carbohydrate (D12451, Research Diets, New Brunswick, NJ, USA). After 8-week HFD feeding, rats were injected intravenously via tail vein with a single low dose of STZ (35 mg/kg) (Sigma). One week after STZ injection, rats showing hyperglycemia (blood glucose ≥15 mmol/L [270 mg/dL]) were considered diabetic.

hPSC-CMs

iCell hiPSC-CMs with >95% purity (CMC-100-110-001) were purchased from Fujifilm Cellular Dynamics, Inc. (Madison, WI) and used as recommended by the manufacturer. This cell line was generated from fibroblasts obtained from female donor (<18 years old) via retroviral transduction.

MV isolation

MV fragments were isolated from the epididymal fat of male Sprague-Dawley (retired breeders, Charles River) or GFP rats (SD-Tg(UBC-EGFP)2BalRrrc; RRRC) or from gonadal and visceral fat of female GFP rats by limited collagenase digestion (type I collagenase, Worthington Biochemical Company, NJ) and selective size sieving as previously described (Hoying et al., 1996; Sun et al., 2020). MVs were prepared at a density of 50,000 MVs/mL in 6 mg/mL type I collagen (BD Biosciences).

MI and cell implantation

Male RNU rats (age 7–8 weeks, Charles River) weighing 180–200 g were used. MI was generated by the permanent ligation of the LCA as previously described (Mihic et al., 2015; Sun et al., 2020). In brief, rats were anesthetized with isoflurane, intubated, and mechanically ventilated with room air supplemented with oxygen. The heart was exposed by an open thoracotomy and subjected to permanent ligation of the left anterior descending artery by 7-0 prolene suture. Two weeks after LCA, the rats meeting the echocardiographic inclusion criterion (FS 20%–35%) were randomly divided into three groups: MI control (n = 6) and those receiving intracardiac injections of hiPSC-CMs only (n = 8) or with freshly isolated MVs (n = 11). Each rat received an intramuscular injection of 10 × 106 hiPSC-CMs only (Laflamme et al., 2007) (Fujifilm Cellular Dynamics, Inc., Madison, WI) or with 5,000 MVs in 100 μL of 6 mg/mL collagen (BD) using a Hamilton syringe and a 28-gauge needle (Hamilton Company, Reno, NV). The surviving rats were analyzed by echocardiography (details are in the following paragraph). PV loop assessments were done before animals were euthanized at week 4 post implantation. To detect blood perfusion, rats were injected intravenously with 1 mg biotinylated Griffonia Simplicifolia Lectin I (GSL I) (Vector Labs) 10 min before heart dissection. Dissected hearts were fixed in 10% neutral-buffered formalin (Sigma) and processed for histological analysis.

Cardiac function

Echocardiography was performed before MI (pre-ligation baseline), before implantation (day 0), and at week 2 and 4 post implantation. LVIDd and LVIDs were measured using a GE Vivid 7 Dimension with a 10S (10 MHz) pediatric probe, and FS was calculated by the equation: FS = 100 × (LVIDd – LVIDs)/LVIDd (Mihic et al., 2015). Rats with FS < 20% or >35% were excluded from the study. LVIDd and LVIDs are expressed in mm.

Four weeks after implantation, cardiac pressure and volume were measured in triplicate with a conductance catheter (SPR-838, Millar Instruments, Inc., Houston, TX), and real-time PV loops were created at baseline and during vena cava occlusion (Mihic et al., 2015). All PV loop data were analyzed using a cardiac PV analysis program (MPVS-400, Millar Instruments). The ESV, EDV, ESP, end-diastolic pressure, EF, max dP/dt, and min dP/dt were calculated. The time constant of isovolumic pressure decay (tau) was calculated using Weiss method (Regression of log (pressure) versus time). Investigators performing cardiac functional analysis were blinded to treatment groups.

Histology and immunocytochemistry

Formalin-fixed hearts were cut into uniform 2-mm-thick transverse slices from apex to base using a matrix slicer (Zivic Instruments, Pittsburgh, PA) before standard paraffin embedding. Each heart usually could be cut into 4–5 slides, which all could be embedded together and then whole mounted on one 25 mm × 75 mm glass slide. 5-μm serial sections were cut, and every 10th slide was stained with hematoxylin-eosin (Sigma) or picrosirius red (Abcam) for quantification of infarct size, or stained for human-specific Ku80 and cTnT or MHC to identify the distribution of the graft. Immunostaining was performed with antibodies directed against human specific Ku80 (1:300, Cell signaling), cTnT (1:100, Thermo Fisher Scientific), sarcomeric α-actinin (1:100, clone EA-53, Sigma), sarcomeric MHC (1:20; clone MF-20, Developmental Studies Hybridoma Bank), Ki67 (1:200, BD Biosciences), CD31/PECAM (1:200, Novus), and GFP (1:1000, Novus). For immunofluorescence staining, Alexa 488 or 568-conjugated secondary antibodies or streptavidin (Thermo Fisher Scientific) were used. For bright-field detection, biotinylated goat anti-mouse or rabbit secondary antibodies (Vector Labs) were used in conjunction with the Vectastain Avidin/Biotin Complex (ABC) Kit (Vector labs) followed by HRP/DAB or alkaline phosphatase/Vector Red (Vector Labs, Burlingame, CA, USA). Cells and sections were counterstained with Hoechst 33342 (1:1,000, Sigma) or hematoxylin (Sigma) to detect nuclei. Sections were mounted using Fisher Chemical permount mounting medium (Fisher Scientific) or vectashield antifade mounting medium (Vector labs). Confocal fluorescence images were captured using Olympus Fluoview 1000 laser scanning confocal microscope.

Images of histological microscope slides were digitally captured by a Leica AT2 Scanscope for morphometric analyses using ImageJ (1.49v). Infarct size was expressed as the percentage of infarct area over total left-ventricular cross-sectional area. Infarct border zone thickness was measured between the infarct and remote (normal) area. Collagen-positive area was determined by the amount of red in picrosirius red staining. Cell surface area was measured by WGA staining using ImageJ. All quantification of histologic parameters was done in a blinded manner.

Statistics

We used ANOVA for analysis involving three groups. For analysis of time-course changes, a paired t test analysis of means was used for samples with normal distribution. hiPSC-CM survival was analyzed by Shapiro-Wilk test followed by Mann-Whitney rank-sum test. All animals were individually coded and maintained until all data were acquired and analyzed. Values are expressed as mean ± SEM unless otherwise stated. p < 0.05 was considered statistically significant.

Cytokine array

Isolated MVs were prepared at a density of 20,000 MVs/mL in 3 mg/mL type I collagen (BD Biosciences) and cultured in DMEM (Thermo Fisher Scientific) containing 10% FBS (Wisent Bioproducts). A collagen-only construct without MV was incubated with same media as control. 4 days later, 500 μL of MV culture or control media were taken for the cytokine array as per manufacturer’s instruction (R&D Systems). The membrane was developed, and exposure was collected using MicroChemi (v.4.2, DNR Bio Imaging System). The pixel densities were analyzed using ImageJ (1.49v).

Antibody-mediated IL-6 depletion

Isolated MVs were resuspended in collagen gel and cultured in media for 4 days. Then the media were collected. IL-6 antibody (1:150, Thermo Fisher Scientific) or rabbit IgG isotype control antibody was added and incubated at 4°C overnight. After addition of Protein A/G Sepharose beads (Thermo Fisher Scientific), the samples were incubated for 2 h at 4°C. Samples were then centrifuged at 1,000 rpm for 5 min, and the supernatant was carefully harvested. Centrifugation at same speed and duration was repeated to ensure complete removal of residual agarose beads. The immunodepletion of IL-6 was confirmed by ELISA as per manufacturer’s instruction (Sigma).

In vitro CM hypertrophy assay

hESC-CMs (ESI-017, day 22 of differentiation) were plated in RPMI media with B27 supplement (Thermo Fisher Scientific), then after 48 h were incubated with the media containing 0%, 25%, 50%, or 75% of MV culture-conditioned media, IL-6-depleted media, or control media for 1 week prior to fixation and immunostaining with alpha-actinin (1:200, Sigma) or cTnT (1:100, Thermo Fisher Scientific) and Hoechst (1:1,000).

Resource availability

Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the corresponding author, Sara S Nunes (sara.vasconcelos@utoronto.ca).

Materials availability

This study did not generate new unique reagents.

Data and code availability

  • •

    This study did not generate new datasets.

  • •

    Raw data and images are available upon request to the corresponding author.

Acknowledgments

This work was supported by grants from the Canadian Institutes of Health Research (PJT153160 and PJT180641) to S.S.N. who holds the John Kitson McIvor Endowed Chair in Diabetes Research.

Declaration of interests

The authors declare no competing interests.

Published: January 23, 2025

Footnotes

Supplemental information can be found online at https://doi.org/10.1016/j.stemcr.2024.102394.

Supplemental information

Document S1. Figures S1–S6
mmc1.pdf (1.3MB, pdf)
Document S2. Article plus supplemental information
mmc2.pdf (5.4MB, pdf)

References

  1. Aghazadeh Y., Poon F., Sarangi F., Wong F.T.M., Khan S.T., Sun X., Hatkar R., Cox B.J., Nunes S.S., Nostro M.C. Microvessels support engraftment and functionality of human islets and hESC-derived pancreatic progenitors in diabetes models. Cell Stem Cell. 2021;28:1936–1949.e8. doi: 10.1016/j.stem.2021.08.001. [DOI] [PubMed] [Google Scholar]
  2. Allard J., Li K., Lopez X.M., Blanchard S., Barbot P., Rorive S., Decaestecker C., Pochet R., Bohl D., Lepore A.C., et al. Immunohistochemical toolkit for tracking and quantifying xenotransplanted human stem cells. Regen. Med. 2014;9:437–452. doi: 10.2217/rme.14.26. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Altalhi W., Hatkar R., Hoying J.B., Aghazadeh Y., Nunes S.S. Type I Diabetes Delays Perfusion and Engraftment of 3D Constructs by Impinging on Angiogenesis; Which can be Rescued by Hepatocyte Growth Factor Supplementation. Cell. Mol. Bioeng. 2019;12:443–454. doi: 10.1007/s12195-019-00574-3. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Altalhi W., Sun X., Sivak J.M., Husain M., Nunes S.S. Diabetes impairs arterio-venous specification in engineered vascular tissues in a perivascular cell recruitment-dependent manner. Biomaterials. 2017;119:23–32. doi: 10.1016/j.biomaterials.2016.12.003. [DOI] [PubMed] [Google Scholar]
  5. Beckerman M., Harel C., Michael I., Klip A., Bilan P.J., Gallagher E.J., LeRoith D., Lewis E.C., Karnieli E., Levenberg S. GLUT4-overexpressing engineered muscle constructs as a therapeutic platform to normalize glycemia in diabetic mice. Sci. Adv. 2021;7 doi: 10.1126/sciadv.abg3947. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Bruin J.E., Saber N., Braun N., Fox J.K., Mojibian M., Asadi A., Drohan C., O'Dwyer S., Rosman-Balzer D.S., Swiss V.A., et al. Treating diet-induced diabetes and obesity with human embryonic stem cell-derived pancreatic progenitor cells and antidiabetic drugs. Stem Cell Rep. 2015;4:605–620. doi: 10.1016/j.stemcr.2015.02.011. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Chatterjee S., Khunti K., Davies M.J. Type 2 diabetes. Lancet. 2017;389:2239–2251. doi: 10.1016/S0140-6736(17)30058-2. [DOI] [PubMed] [Google Scholar]
  8. Chong J.J.H., Yang X., Don C.W., Minami E., Liu Y.W., Weyers J.J., Mahoney W.M., Van Biber B., Cook S.M., Palpant N.J., et al. Human embryonic-stem-cell-derived cardiomyocytes regenerate non-human primate hearts. Nature. 2014;510:273–277. doi: 10.1038/nature13233. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Dhahri W., Sadikov Valdman T., Wilkinson D., Pereira E., Ceylan E., Andharia N., Qiang B., Masoudpour H., Wulkan F., Quesnel E., et al. In Vitro Matured Human Pluripotent Stem Cell-Derived Cardiomyocytes Form Grafts With Enhanced Structure and Function in Injured Hearts. Circulation. 2022;145:1412–1426. doi: 10.1161/CIRCULATIONAHA.121.053563. [DOI] [PubMed] [Google Scholar]
  10. Festing M.F., May D., Connors T.A., Lovell D., Sparrow S. An athymic nude mutation in the rat. Nature. 1978;274:365–366. doi: 10.1038/274365a0. [DOI] [PubMed] [Google Scholar]
  11. Glass C.E., Singal P.K., Singla D.K. Stem cells in the diabetic infarcted heart. Heart Fail. Rev. 2010;15:581–588. doi: 10.1007/s10741-010-9172-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Gomez-Garcia M.J., Quesnel E., Al-Attar R., Laskary A.R., Laflamme M.A. Maturation of human pluripotent stem cell derived cardiomyocytes in vitro and in vivo. Semin. Cell Dev. Biol. 2021;118:163–171. doi: 10.1016/j.semcdb.2021.05.022. [DOI] [PubMed] [Google Scholar]
  13. Hinkel R., Howe A., Renner S., Ng J., Lee S., Klett K., Kaczmarek V., Moretti A., Laugwitz K.L., Skroblin P., et al. Diabetes Mellitus-Induced Microvascular Destabilization in the Myocardium. J. Am. Coll. Cardiol. 2017;69:131–143. doi: 10.1016/j.jacc.2016.10.058. [DOI] [PubMed] [Google Scholar]
  14. Hirota H., Izumi M., Hamaguchi T., Sugiyama S., Murakami E., Kunisada K., Fujio Y., Oshima Y., Nakaoka Y., Yamauchi-Takihara K. Circulating interleukin-6 family cytokines and their receptors in patients with congestive heart failure. Heart Ves. 2004;19:237–241. doi: 10.1007/s00380-004-0770-z. [DOI] [PubMed] [Google Scholar]
  15. Hirota H., Yoshida K., Kishimoto T., Taga T. Continuous activation of gp130, a signal-transducing receptor component for interleukin 6-related cytokines, causes myocardial hypertrophy in mice. Proc. Natl. Acad. Sci. USA. 1995;92:4862–4866. doi: 10.1073/pnas.92.11.4862. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Hoying J.B., Boswell C.A., Williams S.K. Angiogenic potential of microvessel fragments established in three-dimensional collagen gels. In vitro cellular & developmental biology. Animal. 1996;32:409–419. doi: 10.1007/BF02723003. [DOI] [PubMed] [Google Scholar]
  17. Hu S.H., Jiang T., Yang S.S., Yang Y. Pioglitazone ameliorates intracerebral insulin resistance and tau-protein hyperphosphorylation in rats with type 2 diabetes. Exp. Clin. Endocrinol. Diabetes. 2013;121:220–224. doi: 10.1055/s-0032-1333277. [DOI] [PubMed] [Google Scholar]
  18. Jia G., Hill M.A., Sowers J.R. Diabetic Cardiomyopathy: An Update of Mechanisms Contributing to This Clinical Entity. Circ. Res. 2018;122:624–638. doi: 10.1161/CIRCRESAHA.117.311586. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Laflamme M.A., Chen K.Y., Naumova A.V., Muskheli V., Fugate J.A., Dupras S.K., Reinecke H., Xu C., Hassanipour M., Police S., et al. Cardiomyocytes derived from human embryonic stem cells in pro-survival factors enhance function of infarcted rat hearts. Nat. Biotechnol. 2007;25:1015–1024. doi: 10.1038/nbt1327. [DOI] [PubMed] [Google Scholar]
  20. Loai S., Zhou Y.Q., Vollett K.D.W., Cheng H.L.M. Skeletal Muscle Microvascular Dysfunction Manifests Early in Diabetic Cardiomyopathy. Front. Cardiovasc. Med. 2021;8 doi: 10.3389/fcvm.2021.715400. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Mihic A., Cui Z., Wu J., Vlacic G., Miyagi Y., Li S.H., Lu S., Sung H.W., Weisel R.D., Li R.K. A Conductive Polymer Hydrogel Supports Cell Electrical Signaling and Improves Cardiac Function After Implantation into Myocardial Infarct. Circulation. 2015;132:772–784. doi: 10.1161/CIRCULATIONAHA.114.014937. [DOI] [PubMed] [Google Scholar]
  22. Nalbach L., Roma L.P., Schmitt B.M., Becker V., Körbel C., Wrublewsky S., Pack M., Später T., Metzger W., Menger M.M., et al. Improvement of islet transplantation by the fusion of islet cells with functional blood vessels. EMBO Mol. Med. 2021;13 doi: 10.15252/emmm.202012616. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Plenz G., Eschert H., Erren M., Wichter T., Böhm M., Flesch M., Scheld H.H., Deng M.C. The interleukin-6/interleukin-6-receptor system is activated in donor hearts. J. Am. Coll. Cardiol. 2002;39:1508–1512. doi: 10.1016/s0735-1097(02)01791-6. [DOI] [PubMed] [Google Scholar]
  24. Protze S.I., Lee J.H., Keller G.M. Human Pluripotent Stem Cell-Derived Cardiovascular Cells: From Developmental Biology to Therapeutic Applications. Cell Stem Cell. 2019;25:311–327. doi: 10.1016/j.stem.2019.07.010. [DOI] [PubMed] [Google Scholar]
  25. Reed M.J., Meszaros K., Entes L.J., Claypool M.D., Pinkett J.G., Gadbois T.M., Reaven G.M. A new rat model of type 2 diabetes: the fat-fed, streptozotocin-treated rat. Metabolism. 2000;49:1390–1394. doi: 10.1053/meta.2000.17721. [DOI] [PubMed] [Google Scholar]
  26. Rolstad B. The athymic nude rat: an animal experimental model to reveal novel aspects of innate immune responses? Immunol. Rev. 2001;184:136–144. doi: 10.1034/j.1600-065x.2001.1840113.x. [DOI] [PubMed] [Google Scholar]
  27. Saeedi P., Petersohn I., Salpea P., Malanda B., Karuranga S., Unwin N., Colagiuri S., Guariguata L., Motala A.A., Ogurtsova K., et al. Global and regional diabetes prevalence estimates for 2019 and projections for 2030 and 2045: Results from the International Diabetes Federation Diabetes Atlas, 9(th) edition. Diabetes Res. Clin. Pract. 2019;157 doi: 10.1016/j.diabres.2019.107843. [DOI] [PubMed] [Google Scholar]
  28. Sahin K., Onderci M., Tuzcu M., Ustundag B., Cikim G., Ozercan I.H., Sriramoju V., Juturu V., Komorowski J.R. Effect of chromium on carbohydrate and lipid metabolism in a rat model of type 2 diabetes mellitus: the fat-fed, streptozotocin-treated rat. Metabolism. 2007;56:1233–1240. doi: 10.1016/j.metabol.2007.04.021. [DOI] [PubMed] [Google Scholar]
  29. Schmidt-Arras D., Rose-John S. IL-6 pathway in the liver: From physiopathology to therapy. J. Hepatol. 2016;64:1403–1415. doi: 10.1016/j.jhep.2016.02.004. [DOI] [PubMed] [Google Scholar]
  30. Shiba Y., Fernandes S., Zhu W.Z., Filice D., Muskheli V., Kim J., Palpant N.J., Gantz J., Moyes K.W., Reinecke H., et al. Human ES-cell-derived cardiomyocytes electrically couple and suppress arrhythmias in injured hearts. Nature. 2012;489:322–325. doi: 10.1038/nature11317. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Si Y., Zhao Y., Hao H., Liu J., Guo Y., Mu Y., Shen J., Cheng Y., Fu X., Han W. Infusion of mesenchymal stem cells ameliorates hyperglycemia in type 2 diabetic rats: identification of a novel role in improving insulin sensitivity. Diabetes. 2012;61:1616–1625. doi: 10.2337/db11-1141. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Srinivasan K., Viswanad B., Asrat L., Kaul C.L., Ramarao P. Combination of high-fat diet-fed and low-dose streptozotocin-treated rat: a model for type 2 diabetes and pharmacological screening. Pharmacol. Res. 2005;52:313–320. doi: 10.1016/j.phrs.2005.05.004. [DOI] [PubMed] [Google Scholar]
  33. Strobel H.A., Gerton T., Hoying J.B. Vascularized adipocyte organoid model using isolated human microvessel fragments. Biofabrication. 2021;13 doi: 10.1088/1758-5090/abe187. [DOI] [PubMed] [Google Scholar]
  34. Sun X., Wu J., Qiang B., Romagnuolo R., Gagliardi M., Keller G., Laflamme M.A., Li R.K., Nunes S.S. Transplanted microvessels improve pluripotent stem cell-derived cardiomyocyte engraftment and cardiac function after infarction in rats. Sci. Transl. Med. 2020;12 doi: 10.1126/scitranslmed.aax2992. [DOI] [PubMed] [Google Scholar]
  35. Szkudelski T. The mechanism of alloxan and streptozotocin action in B cells of the rat pancreas. Physiol. Res. 2001;50:537–546. [PubMed] [Google Scholar]
  36. Tan Y., Zhang Z., Zheng C., Wintergerst K.A., Keller B.B., Cai L. Mechanisms of diabetic cardiomyopathy and potential therapeutic strategies: preclinical and clinical evidence. Nat. Rev. Cardiol. 2020;17:585–607. doi: 10.1038/s41569-020-0339-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Torre-Amione G., Kapadia S., Lee J., Bies R.D., Lebovitz R., Mann D.L. Expression and functional significance of tumor necrosis factor receptors in human myocardium. Circulation. 1995;92:1487–1493. doi: 10.1161/01.cir.92.6.1487. [DOI] [PubMed] [Google Scholar]
  38. Yin F.C., Spurgeon H.A., Rakusan K., Weisfeldt M.L., Lakatta E.G. Use of tibial length to quantify cardiac hypertrophy: application in the aging rat. Am. J. Physiol. 1982;243:H941–H947. doi: 10.1152/ajpheart.1982.243.6.H941. [DOI] [PubMed] [Google Scholar]
  39. Zhang M., Lv X.Y., Li J., Xu Z.G., Chen L. The characterization of high-fat diet and multiple low-dose streptozotocin induced type 2 diabetes rat model. Exp. Diabetes Res. 2008;2008 doi: 10.1155/2008/704045. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Zhou T., Hu Z., Yang S., Sun L., Yu Z., Wang G. Role of Adaptive and Innate Immunity in Type 2 Diabetes Mellitus. J. Diabetes Res. 2018;2018 doi: 10.1155/2018/7457269. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Document S1. Figures S1–S6
mmc1.pdf (1.3MB, pdf)
Document S2. Article plus supplemental information
mmc2.pdf (5.4MB, pdf)

Data Availability Statement

  • •

    This study did not generate new datasets.

  • •

    Raw data and images are available upon request to the corresponding author.

Articles from Stem Cell Reports are provided here courtesy of Elsevier

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