Angiotensin-converting enzyme 2 (ACE2) is a transmembrane protein consisting of a C-terminal region, a membrane anchoring region, and an N-terminal extracellular region containing the catalytic domain and a signal peptide. The catalytic domain is a single HEMGH zinc-binding sequence [1, 2]. Through this catalytic domain, ACE2 exerts its monocarboxipeptidase actions on a single aminoacid of Angiotensin (ANG)-II C-terminal end, promoting Ang-(1-7) production, and counterbalancing ACE/Ang-II axis [3]. Ang-(1-7) triggers its actions through MAS G-protein-coupled receptor, opposing ACE/ANG-II axis, which include vasodilatation and vascular protective actions as well as anti-fibrotic, anti-proliferative, and anti-inflammatory effects [4, 5].

ACE2 has been proposed as a therapeutic target for being a negative regulator of the renin–angiotensin system (RAS) [6, 7], thus, opening up new wide range of possibilities in the treatment of renal and cardiovascular diseases. Several studies have been performed in different organs and pathologies, mainly focused on kidney and heart diseases [8,9,10,11]. Previous studies based on pharmacologic inhibition [12, 13] and genetic ablation [14,15,16] constituted a key tool within the knowledge of ACE2 in renal and heart pathophysiology. ACE2 has been shown to be involved in cardiovascular and kidney disease. In the nonobese diabetic (NOD) mouse, we previously demonstrated that circulating and renal ACE2 is altered and the administration of insulin restores this alteration coupled with normalization of renal parameters [17]. NOD mice spontaneously develop an insulin-dependent diabetes mellitus (DM) similarly to type 1 DM in humans including hyperglycemia, glycosuria, hypercholesterolemia, ketonuria, polydipsia, polyuria, and polyphagia [18]. ACE2 is expressed in pancreas and increased in different models of diabetes, such as NOD (type 1) and db/db mice (type 2) [8, 19, 20]. The effect of ACE2 deletion or overexpression on pancreas from diabetic mice has only been tested in type 2 diabetes [21, 22]. To better understand the contribution of ACE2 in type 1 diabetes, we study the effects of global ACE2 deficiency on pancreatic function and on diabetic kidney disease progression in the spontaneous model of type 1 diabetes, the NOD mice. This model may help to understand (i) the contribution of ACE2 in diabetes onset in this mouse strain and (ii) the role of ACE2 in the progression of diabetic nephropathy in this experimental mouse model (Fig. 1S). Furthermore, the direct effect of ACE2 deletion on proximal renal tubular cell has not been completely studied. We hypothesized that ACE2 deletion exerts a harmful effect on the glucose homeostasis and worsens DN progression in the NOD mice. These effects would be mediated by RAS modulation and by fibrotic processes accompanied by increases in oxidative stress and inflammatory mediators.

Materials and methods


NOD/ShiLtJ (Stock No: 001976) and NOR/LtJ (Stock No: 002050) mice were crossed with C57BL/6 ACE2 knockout (ACE2KO) animals obtained from Dr SB Gurley. The generation of ACE2KO mice was previously published [16, 23]. In this study, the background was changed assuming that after ten backcross generations 99.9% of the genome of the donor background has been successfully transferred onto the receptor strain [24]. After that, NOD.ACE2−/− and NOR.ACE2/− were generated. Only female mice were used because of the higher incidence of diabetes described in them as compared with male mice [17]. Diabetic NOD mice are extensively used to study type 1 diabetes, and the nondiabetic NOR mice, which carry a portion of the NOD/ShiLtJ genome are used as control.

Mice were housed in ventilated microisolators under SPF conditions, in a 12-h light–dark cycle with free access to chow and tap water. All procedures were approved by the Barcelona Biomedical Research Park Ethical Committee for Animal Experimentation and Catalan Government (MSO-12-1447; DAAM 6617).

At the end of the follow-up, mice were anesthetized by intraperitoneal (i.p.) injection of sodium pentobarbital, and blood was collected by intracardiac puncture. Mice were perfused with cold phosphate buffer solution prior tissue removal. The left kidney and half of the right kidney were snap frozen with liquid nitrogen and kept at −80 °C for further analysis. Half of the right kidney and pancreas were maintained in formalin solution, neutral buffered 10% (Sigma), and paraffin embedded for histological studies.

Cumulative diabetes incidence was tested in NOD.ACE2+/+ (n = 107) and NOD.ACE2/− (n = 124). From 12 weeks of age, blood glucose (BG) levels from tail vein were measured every 4 weeks till 24 weeks of age. A mouse was considered diabetic when BG levels exceeded 250 mg/dl for two consecutive days. At the age of 26 weeks, all remaining mice were killed by carbon dioxide overdose.

ACE2 deletion in pancreas and glucose homeostasis

I.p. glucose tolerance test was performed in NOD.ACE2+/+ (n = 107) and NOD.ACE2−/ (n = 124) at 12, 16, 20, and 24 weeks of age, as previously described [21]. Female mice were fasted for 12 h and weighted. D-glucose (Sigma-Aldrich, USA) bolus was injected i.p. at 2 g/Kg body weight. Blood samples from the tail vein were obtained at 0, 15, 30, 60, and 120 min after injection, and BG levels were measured with the ACCU-CHEK Compact meter system (Roche Diabetes Care, Spain). Results were expressed as area under curve (AUC) between 0 and 60 min after injection, and as BG levels between 0 and 120 min. Plasma insulin concentrations at 0 and 5 min after glucose injection were measured with the Ultra-Sensitive Mouse Insulin kit (Crystal Chem Inc., Netherlands) according to manufacturer instructions. AUC was calculated between 0 and 5 min.

Insulin tolerance test was performed 24 h after the glucose tolerance test. Animals were fasted for 5 h and weighted. An insulin dose of 0.75 U/Kg body weight (Humulin R, Eli Lilly) was i.p. injected. BG levels from the mouse tail vein were measured at 0, 15, 30, 60, and 120 min after insulin bolus injection. Results were expressed as BG levels from 0 to 120 min [21].

Paraffin-embedded pancreas from 12-week NOD.ACE2/− (n = 7) and their respective NOD wild-type mice (n = 7) were cut into 3-µm sections, deparaffinized in xylene, and rehydrated through graded alcohols. Primary antibodies for insulin (1:1000; Cell Signaling), ACE (1:500; Bioworld), AT1R (1:500; Santa Cruz Biotechnology), the oxidative stress marker nitrotyrosine (1:1000; Merck Millipore), and the key mediator of necroptosis RIPK1 (1:500; Cell Signaling) were used for the immunohistochemistry techniques. EnVision+ System- HRP Labeled Polymer Anti-Rabbit (Dako) and mouse anti-goat IgG-HRP (Santa Cruz Biotechtechnology) were used as secondary antibodies. Samples were counterstained with hematoxylin. All analyses were performed in a blinded fashion. Islet staining was quantified by ImageJ Software 1.47v. Briefly, at minimum of ten islets from pancreatic sections were measured at ×200 of magnification. Specific brown signal was digitally isolated, quantified, and expressed as mean gray value per μm2 (ImageJ, Software 1.47v, NIH).

ACE2 deletion in kidney and diabetic nephropathy progression

For the study of the diabetic nephropathy progression, NOD.ACE2+/+ (n = 13) and NOR.ACE2+/+ (n = 13) and NOD.ACE2/− (n = 10) and NOR.ACE2−/− (n = 10) animals were followed starting at 12 weeks of age. BG levels from tail vein were measured after 4-h fasting. Mice showing two consecutive measures higher than 250 mg/dl were considered diabetic. Diabetic mice were studied for 30 days following the diabetes diagnosis. Age ranged between 16 and 24 weeks.

Systolic and diastolic blood pressure (SBP and DBP) were measured throughout the last week of follow-up using the noninvasive tail-cuff method CODA (Kent Scientific Corporation, Torrington, Connecticut, USA). Animals were previously habituated and trained for valid measurements [17, 23, 25]. SBP and DBP were expressed in mmHg.

After 30 days of follow-up and before end-point surgery, glomerular filtration rate (GFR) was tested by plasmatic clearance of a single-bolus injection of inulin-FITC (Sigma-Aldrich, USA). Conscious animals previously weighted and immobilized were injected with 2.74 µL/gr of body weight of 5% dialyzed inulin-FITC via caudal vein. Blood was extracted at 3, 7, 10, 15, 35, 55, and 75 min after injection and collected in lithium heparinized capillary tubes (Microvette® 100, Starstedt). GFR values were expressed as ml/min per g of body weight as previously published [23, 26, 27].

Spot urine obtained at the end of the study was used for determining albuminuria-to-creatinine ratio (ACR). Urinary albumin and creatinine levels were measured by ELISA (Albuwell M; Exocell, Philadelphia, Pennsylvania, USA) and colorimetric assay (Creatinine Companion; Exocell), respectively, following manufacturer’s instructions. ACR was expressed as µgAlb/mgCrea.

Kidney histology was analyzed on 3-μm sections from tissue embedded in paraffin blocks. After deparaffination and rehydration, sections were stained with periodic acid-Schiff for assessment of mesangial and glomerular areas by pixel counts on a minimum of 20 glomeruli per section, under ×400 magnification (Image J, NIH). Podocyte counts were determined by staining for WT-1. For this purpose, antigen retrieval was performed in pressure cooker with citrate buffer, pH6. After peroxidase activity blocking with 3% H2O2, sections were incubated with rabbit anti-WT-1 (1:1000, Santa Cruz Biotechnology). At the end of the process, 20 microphotographs of glomeruli were taken at ×400 for each animal. WT-1 positive nuclei per glomeruli were double blinded counted and referred to total cell number [17, 28].

Angiotensin-converting enzyme (ACE) activity was assayed in serum and renal cortex using a fluorometric method [28]. Two microliters of serum or 0.5 µg of renal cortex protein was incubated in duplicate with 73 µL of reaction buffer (0.5 M borate buffer and 5.45 M N-hippuryl-His-Leu (Sigma-Aldrich, USA)) for 15 or 25 min at 37 oC, respectively. Twenty milligrams per milliliter of o-Phthalaldehyde (Sigma-Aldrich, USA) was added to the reaction and formed a fluorescent adduct with the ACE-catalyzed product L-histidyl-L-leucine. Fluorescence was measured at λex = 360 nm and λem = 485 nm. ACE enzymatic activity in serum and renal cortex was expressed as RFU/µL/min and RFU/µg/min, respectively.

Changes in gene expression of renal RAS components were analyzed. For this purpose, total RNA was extracted from 50 mg of renal cortex using Tripure isolation Reagent (Roche, Germany). RNA quantity and purity were analyzed with NanoDrop ND-1000. cDNA was synthesized from 1 µg of RNA using the High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems, USA). Real time PCR was performed on Light Cycler 480 System (Roche, Germany) using SYBR Green Master Mix 2× (Roche, Germany). Primers are shown in Table 1. Gapdh or Hprt were the housekeeping genes depending on sample type. The ΔΔCt method was used to calculate the relative quantity of the gene expression.

Table 1 Primer list used for gene expression analysis.

ACE2 deletion in renal proximal tubular cells and gene expression modifications

Isolated primary cell culture was used to better delineate the role of ACE2 in the proximal tubular cell. For this purpose, twelve-week-old NOD.ACE2−/− and their respective NOD wild-type mice were anesthetized and kidneys removed. The tissue was kept in ice-cold HBSS containing penicillin 100 IU/ml and streptomycin 100 μg/ml (P/S) before starting the digestion protocol following a modification from the previously published by Terryn et al. [29]. Briefly, renal cortices in HBSS-P/S were dissected in 1-mm-wide fragments and digested with 0.5 mg/mL type 2 collagenase solution at 37 °C for 30 min. Then, tissue was mushed on two nylon sieves (pore size 200 and 80 μm, consecutively) considering that proximal tubules (PT) were retained on the 80-μm sieve. After several washes with HBSS-P/S-1% FBS, tubules were retrieved by flushing the sieve in the reverse direction. The solution was centrifuged for 20 min at 120 g, washed, and then resuspended in culture medium (DMEM/F12 supplemented with 10% heat-inactivated FBS, 2 mmol/l L-glutamine, 40 ng/ml hydrocortisone, 1% ITS, 10 ml/l NEAA and P/S). The PT fragments were grown on 1% gelatin-coated Petri dishes in 95% air—5% CO2 in a standard humidified incubator for 8–10 days until they reached 60–80% confluency. Medium was changed on day 2 and then every 6 days. Trypsin-EDTA was used for subculturing after passage 4. Previously, TrypLE™ Express Enzyme (Gibco) was used as dissociating method. Cells were characterized by gene expression after RNA extraction as described above. Primers used are listed in Table 1.

Statistical analysis

Quantitative variables are expressed as mean ± standard error of the mean. Variable distribution was assessed according the Kolmogorov–Smirnov test. Mean comparison was made using t Student and U Mann–Whitney tests when variables were not following a normal distribution. For comparing means throughout the 12 weeks of follow-up, the variance of the results was studied using the Kruskal–Wallis test. Cumulative diabetes incidence was calculated using the Kaplan–Meier estimation, and statistical significance was evaluated by the log-rank test. Statistically significance was stablished when p ≤ 0.05. SPSS version 18.0 for Windows (SPSS Inc. Chicago, USA) was used for analysis.


Effect of ACE2 deletion in pancreas and glucose homeostasis

Body weight

NOD.ACE2/− animals showed lower body weight as compared with NOD.ACE2+/+ at 16, 20, and 24 weeks of age (Table 2). NOR.ACE2−/− presented lower body weight as compared with NOR.ACE2+/+ at 12 and 16 weeks of age (Table 2). In addition, among KO mice, body weight was lower in NOD.ACE2−/− animals as compared with NOR.ACE2−/− at 24 weeks of age.

Table 2 Body weight after 12-h fasting at 12, 16, 20, and 24 weeks of age in NOD.

Diabetes incidence

The percentage of diabetic mice at the end of the study was 43.9% for NOD.ACE2+/+ (n = 107) and 55.6% for NOD.ACE2−/− (n = 124). The median diabetes onset time was 25 weeks of age (interquartile range: 23–25) for the wild-type mice and 24 weeks (interquartile range: 21–25) for the knockout mice. Although ACE2 deletion seems to have higher affected diabetes development, the cumulative diabetes incidence did not reach statistical significance compared with the wild-type mice (p = 0.072) (Fig. 2S).

Glucose tolerance test

NOD.ACE2/− mice presented higher BG levels in response to glucose bolus injection as compared with NOD.ACE2+/+. In NOD.ACE2−/− at 12 weeks of age, BG levels were significantly increased at 15, 30, and 60 min after bolus injection in 12-week-old mice (Fig. 1a). BG levels were also significantly higher in NOD.ACE2/− at 15, and 30 min after glucose bolus injection as compared with NOD.ACE2+/+ in 16-week-old mice (Fig. 1b). AUC between 0 and 60 min after the i.p. glucose test for each age point was calculated (Fig. 1e). NOD.ACE2−/− showed decreased glucose bolus tolerance in a first phase (0–60 min after injection), as compared with NOD.ACE+/+ in 12, and 16-week-old mice.

Fig. 1: Intraperitoneal glucose tolerance tests.
figure 1

Blood glucose curves in NOD.ACE2+/+, NOD.ACE2/−, NOR.ACE2+/+, and NOR.ACE2-/- mice in response to glucose bolus administration over 2 h of follow-up. ad Curves at 12, 16, 20, and 24 weeks of age and e AUC values between 0 and 60 min of the glucose test in NOD.ACE2/− animals compared with NOD.ACE2+/+ at 12, 16, 20, and 24 weeks of age. fi Curves at 12, 16, 20, and 24 weeks of age and j AUC values in NOR.ACE2/− animals compared with NOR.ACE2+/+. *p < 0.05 ACE2−/− vs. ACE2+/+.

In contrast to NOD strains, NOR.ACE2/− presented lower increase in BG levels in response to glucose bolus administration as compared with NOR.ACE2+/+ (Fig. 1f–j). In NOR.ACE2−/−, BG levels were significantly decreased at 15, 30, and 120 min as compared to NOR.ACE2+/+ in 24-week-old mice.

Intraperitoneal insulin tolerance test

After insulin administration, NOD.ACE2−/− presented faster decrease in BG and, therefore, higher sensitivity to insulin injection as compared with NOD.ACE2+/+ at 15, 30, and 60 min in 12-week-old mice. The same pattern was observed at 15, 30, 60, and 120 min in 16-week-old mice and at 30, 60, and 120 min in 20-week-old mice. In older animals, fewer differences were shown, only after 120 min of insulin injection (Fig. 2a–d).

Fig. 2: Insulin tolerance test in NOD and NOR mice.
figure 2

Blood glucose curves (mg/dL) in response to insulin bolus over 2 h after injection. a–d Curves at 12, 16, 20, and 24 weeks of age for NOD mice. e–h Curves at 12, 16, 20, and 24 weeks of age for NOR mice. *p ≤ 0.05 ACE2−/− vs. ACE2+/+.

Insulin release

Insulin secretion was assessed by AUC of insulin levels determined between 0 and 5 min after glucose bolus injection in NOD mice. As depicted in Fig. 3S NOD.ACE2−/− group showed lower insulin secretion as compared with NOD.ACE2+/+ in 12-week-old mice, without reaching statistical significance. This trend was also found on 16-week-old mice.

Histological analysis of pancreas in prediabetic mice

Insulin expression in pancreatic β-cells was studied by immunohistochemistry in 12-week-old NOD.ACE2+/+ and NOD.ACE2−/− mice. ACE2KO mice showed a decreased insulin staining inside the pancreatic islet as compared with NOD.ACE2+/+ (Fig. 3a, b).

Fig. 3: Insulin- and RIPK1-positive areas in β-pancreatic cells in nondiabetic animals at 12 weeks of age.
figure 3

a Quantitative analysis of pancreatic islet insulin expression. Mean area with insulin-positive staining per pancreas in nondiabetic NOD.ACE2−/− vs. NOD.ACE2+/+ mice; b representative β-pancreatic islets with insulin-positive staining in microphotographs taken at ×200 magnification; c representative β-pancreatic islets with RIPK1-positive staining in microphotographs taken at ×200 magnification; d mean of RIPK1-positive area in pancreatic islets of NOD.ACE2−/− and NOD.ACE2+/+ mice. *p ≤ 0.05 NOD.ACE2−/− vs. NOD.ACE2+/+.

TUNEL assay was performed to test changes in pancreatic β-cells related to cell death and apoptosis. No differences were observed between NOD.ACE2−/− and NOD.ACE2+/+ animals (data not shown). Interestingly, RIPK1-positive staining, a regulator of necroptosis, was significantly increased in NOD.ACE2−/− as compared with NOD.ACE2+/+ in 12-week-old mice (Fig. 3c, d). When we wanted to explore insulin expression in pancreatic β-cells from older animals with established diabetes (4 weeks after onset), very few islets were found. Although we observed a trend to higher number of infiltrating cells in NOD.ACE2/− mice, the very low number of pancreatic islets per animal was considered not enough for comparisons.

To evaluate modifications in RAS components, ACE and AT1R were immunolocalized in pancreas (Fig. 4). In NOD.ACE2−/− mice, ACE- and AT1R-positive areas were significantly increased into the pancreatic islet as compared with NOD.ACE2+/+ mice.

Fig. 4: ACE- and AT1R-positive areas in β-pancreatic cells.
figure 4

a Representative β-pancreatic islets with ACE-positive staining in microphotographs taken at ×200 magnification; b mean of ACE-positive area in pancreatic islets of NOD.ACE2−/− and NOD.ACE2+/+ nondiabetic animals at 12 weeks of age; c representative β-pancreatic islets with AT1R-positive staining in microphotographs taken at ×200 magnification; d mean of AT1R-positive area in pancreatic islets of NOD.ACE2−/− and NOD.ACE2+/+ mice. *p ≤ 0.05 NOD.ACE2−/− vs. NOD.ACE2+/+.

As marker of oxidative stress, nitrotyrosine expression was also determined (Fig. 5). NOD.ACE2−/− mice significantly increased nitrotyrosine-islet staining as compared with NOD.ACE2+/+ mice in 12-week-old mice.

Fig. 5: Nitrotyrosine-positive area in β-pancreatic cells.
figure 5

a Representative β-pancreatic islets with nitrotyrosine-positive staining in microphotographs taken at ×200 magnification; b mean of nitrotyrosine-positive area in pancreatic islets of NOD.ACE2−/− and NOD.ACE2+/+ nondiabetic animals at 12 weeks of age. *p ≤ 0.05 NOD.ACE2−/− vs. NOD.ACE2+/+.

Effect of ACE2 deletion in diabetic nephropathy progression after 4 weeks of diabetes

Diabetes, blood pressure, and kidney function

NOD diabetic mice showed higher BG levels than NOR control mice of similar age in both, wild-type and knockout mice. At the end of the study, NOD.ACE2+/+ and NOD.ACE2−/− mice presented significant increase in BG as compared with the baseline levels (Table 3). Diabetic wild-type mice presented decreased body weight as compared with the control NOR mice. In concordance, in NOD.ACE2−/− the body weight was lower as compared with NOR.ACE2−/− (Table 3).

Table 3 Animal characteristics at the beginning and the end of the study after 4 weeks of diabetes in NOD.ACE2+/+, NOD.ACE2−/−, and their respective controls NOR.ACE2+/+, NOR.ACE2−/− mice.

Diabetic mice presented a significant increase in renal and heart weight as compared with nondiabetic mice. The ratio of kidney/body weight was significantly increased in NOD diabetic mice as compared with NOR, in both wild-type and knockout mice. Heart/body weight ratio was decreased in NOD diabetic mice as compared with NOR control mice (Table 3).

SBP was significantly increased in all diabetic NOD mice. However, no differences were observed in DBP between nondiabetic and diabetic mice (Fig. 6a).

Fig. 6: Effect of Ace2 deletion on clinical parameters under NOR and NOD backgrounds.
figure 6

a Systolic and diastolic blood pressure (SBP and DBP) at the end of 30-day follow-up. In black, SBP is shown and in gray, values for DBP are shown; b urinary albumin excretion after 30 days of diabetes; c GFR values using inulin-FITC bolus injection at day 30 of follow-up. Open symbols represent NOR strains and closed symbols, the NOD strains. $p ≤ 0.05 NOR vs. NOD.

Diabetes altered kidney function. ACR was significantly increased in both wild-type and knockout NOD diabetic mice (Fig. 6b). GFR was significantly increased only in NOD.ACE2+/+ mice as compared with NOR.ACE2+/+ mice. Although, NOD.ACE2−/− mice showed similar GFR values as observed in NOR.ACE2/− mice, no statistically differences were found when comparing NOD.ACE2+/+ and NOD.ACE2−/− mice (Fig. 6c).

Renal histology

Mesangial area was significantly increased in diabetic animals as compared with nondiabetic control mice, both in wild-type and knockout mice (Fig. 7b). Podocyte loss was evident in the NOD.ACE2−/− as compared with NOR.ACE2−/− mice. This difference was not seen among wild-type groups (Fig. 7c, d).

Fig. 7: Effect of Ace2 deletion on glomerular characteristics under NOR and NOD backgrounds.
figure 7

a Glomerular area in ACE2/− and ACE2+/+ in NOR and NOD mice; b mesangial area in ACE2/− and ACE2+/+ in NOR and NOD mice; c Podocyte number per glomerulus detected as WT-1-positive cells in ACE2/− and ACE2+/+ in NOR and NOD mice; d representative glomeruli with WT-1-positive cells in microphotographs taken at ×200 magnification. Open symbols represent NOR strains and closed symbols, the NOD strains. $p ≤ 0.05 NOR vs. NOD.

Circulating and renal ACE

Circulating ACE activity was significantly increased in diabetic animals as compared with nondiabetic mice, both in wild-type and knockout mice. In addition, diabetic NOD.ACE2−/− mice showed significant decrease in circulating ACE activity as compared with NOD.ACE2+/+ diabetic animals (Fig. 8a).

Fig. 8: Effect of Ace2 deletion on ACE activity and gene expression under NOR and NOD backgrounds.
figure 8

a Serum ACE enzymatic activity; b renal cortical ACE enzymatic activity; and c ACE gene expression in renal cortex in ACE2−/− and ACE2+/+ in NOR and NOD mice. Open symbols represent NOR strains and closed symbols, the NOD strains. $p ≤ 0.05 NOR vs. NOD; *p ≤ 0.05 ACE2−/− and ACE2+/+.

ACE enzymatic activity was similar among all study groups (Fig. 8b). However, renal ACE gene expression was significantly decreased in NOD.ACE2+/+ diabetic as compared with NOR.ACE2+/+ control mice (Fig. 8c).

Renal expression of other RAS components

AT1R, aminopeptidase A, and neprilysin (NEP) gene expression were significantly decreased in NOD.ACE2+/+ diabetic mice as compared with NOR.ACE2+/+ mice. In contrast, Angiotensin II receptor type 2 (AT2R), and Renin gene expression were significantly increased in diabetes as compared with control wild-type mice. In NOD.ACE2/− mice, diabetes significantly decreased AT1R, NEP, and Chymase gene expression as compared with NOR.ACE2/−. Interestingly, animals with ACE2 deletion showed increased AT2R and Renin gene expression as compared with wild-type control mice (Fig. 9).

Fig. 9: Effect of Ace2 deletion on gene expression of RAS components under NOR and NOD backgrounds.
figure 9

Renal cortex was analyzed for a angiotensinogen; b renin; c neprilysin; d chymase; e apa; f at1r; g at2r; h Masr. Open symbols represent NOR strains and closed symbols, the NOD strains. $p ≤ 0.05 NOR vs. NOD; *p ≤ 0.05 ACE2−/− and ACE2+/+.

Effect of ACE2 deletion on proximal tubular cells

To investigate the influence of ACE2 deletion in renal proximal tubular cells, we analyzed fibrosis and RAS components by quantitative PCR analysis of the mRNA expression. ACE2 knockdown in primary proximal tubular cells was confirmed by gene expression (Fig. 10a). The lack of ACE2 on normal proximal tubular cells induced changes in the expression of genes related to fibrosis and RAS. As shown in Fig. 10b, the lack of ACE2 predisposed to activate fibrosis-related genes in NOD diabetic mice before the diabetes onset. Collagen type 1 gene expression was significantly increased in isolated renal PT cells from NOD.ACE2−/− as compared with isolated renal PT cells from NOD.ACE2+/+ mice.

Fig. 10: Effect of ACE2 deletion on gene expression of RAS components and fibrosis-related genes in isolated primary renal proximal tubular cells under normal glucose conditions.
figure 10

Quantitative PCR analysis of Ace2, Angiotensinogen, Renin, Ace, At2R, Tgfβ, Ctgf, Fn1, and Col1a gene expression in the primary cell cultures. *p ≤ 0.05 NOD.ACE2−/− and NOD.ACE2+/+.


This study demonstrated that the strains generated in our laboratory, NOD.ACE2/− as well as NOR.ACE2/−, are viable, fertile, and lack any gross structural abnormality. At pancreatic level, NOD.ACE2−/− presented decrease in glucose tolerance coupled with decreased insulin expression. The deleterious effect of ACE2 deletion within pancreas from NOD mice was associated with activation of the RAS, increase of RIPK1, and oxidative stress. At renal level, NOD.ACE2/− mice presented reduction in podocyte number without hyperfiltration, suggesting a deleterious effect on renal function. In addition, RAS genes were altered in both the NOD- and the NOR-ACE2KO mice. In normoglycemic conditions, ACE2 deletion in renal proximal tubular cells showed an imbalance in gene expression of RAS components and a predisposition to increase fibrotic genes.

NOD mice is a model that develops spontaneous autoimmune diabetes that mimics the autoimmune or type 1 diabetes in humans, including the presence of pancreatic specific autoantibodies, autoreactive CD4+ and CD8+ T cells, and genetic linkage to disease similar to that found in humans [30]. Previous studies from our group showed that NOD mice represent a useful model to study early renal changes in diabetic nephropathy [17]. In the current study, we demonstrated that animals carrying ACE2 deletion under NOD background lack of any gross structural abnormalities. In concordance, Wong et al. demonstrated that ACE2/yIns2WT/C96Y mice were also viable and fertile [31]. ACE2 loss leads to a decreased animal body weight in the two-studied backgrounds. In agreement, previous studies have confirmed that ACE2 pharmacological inhibition leads to lower body weight. Thus, in STZ-diabetic mice, administration of MLN-4760, a specific synthetic ACE2 inhibitor, decreased body weight as compared with vehicle-treated animals [32, 33]. Body weight decrease in NOR mice were mainly observed after fasting at 12–16 weeks of age. Bernardi et al. also showed a decrease in control mice carrying an ACE2 deletion and, exacerbated when animals were given a high-fat diet [34]. In contrast, other authors do not report body weight differences in ACE2KO mice [35]. These differences could be attributed to different KO generating strategies giving genotypic differences.

In the current study, we showed that ACE2 deletion worsens glucose tolerance in the NOD model. These alterations were evident early, at 12–16 weeks of age, and disappear later in time. In this mouse model, new-onset diabetes develops progressively over weeks (~10%, at 14 weeks and ~20% at 16 weeks of age) and the frequencies are depending on gender (80–90% females and 10–40% males at 30 weeks) [17, 18]. Thus, the observed differences in an early phase of the study indicate an alteration in a phase, which the diabetes onset in this model is less frequent. Studies in diabetic models induced by high-fat diet in ACE2KO mice postulate that the alteration in glucose metabolism may be ascribed in part to changes in glucose transporters such as GLUT-2 or GLUT-4 [35]. However, this alteration may be also associated with marked decrease in insulin expression within the pancreatic β-cells. In this sense, Bindom et al. previously demonstrated that ACE2 overexpression increased islet insulin content in db/db mice above the level observed in db/m mice. In addition, the enhanced insulin content, in db/db mice overexpressing ACE2, was due to enhanced pancreatic β-cell proliferation and reduced apoptosis [21]. Our results are in concordance with these findings showing increased nitrotyrosine accumulation in pancreatic islets of NOD.ACE2−/− mice.

Surprisingly, by TUNEL assay, we did not find an increase in apoptosis-related mechanism in pancreatic β-cell from NOD.ACE2−/−. The absence of differences in apoptosis among KO and WT mice could be related to the presence per se of apoptosis in pancreatic islets of NOD model [36], and once initiated, we are not able to distinguish differences regarding Ace2 gene presence or absence. It is possible that in this model apoptotic events take place in an early phase, as we show in this project an early glucose homeostasis alteration. Unfortunately, it is worth mentioning that the initial approach of this study was not to collect pancreatic samples at 12 weeks of age. Moreover, probably TUNEL assay alone is not the best technique for apoptosis detection because DNA fragments with 3′-OH ends can be produced without apoptosis [37]. Interestingly, RIPK1 was increased in pancreatic islets from NOD.ACE2−/−, implying regulation of necrosis and inflammation [38] as one of the mechanisms that may lead to glucose homeostasis alteration in this animal model.

Furthermore, in vitro studies of pancreatic β-cells have demonstrated the ability of glucose to activate ACE2/Ang-(1-7)/Mas and APN/Ang-IV/IRAP axis [39]. In the same line of evidence Ang-II infusion produces a decrease in ACE2 pancreatic activity and expression accompanied by an increased AT1R expression as well as oxidative stress [40]. Consistently, we now found that ACE2 deletion increases oxidative stress accompanied by an increased ACE as well as AT1R expression in pancreatic islet from NOD mice. The presence of RAS components, ACE and ACE2 have been described in both endocrine and exocrine pancreas in a rat model of obesity type 2 DM [41]. In experimental models of type 2 diabetes, RAS blockade by ARB improved pancreatic β-cell function and glucose tolerance [42]. ACE2 overexpression by gene therapy is associated with decreased BG, improved glucose tolerance, increased insulin secretion, and pancreatic β-cells proliferation and decreased apoptosis through Ang-(1-7) in db/db mice [21]. These results are in line with our results where the opposite effect is observed when Ace2 expression is deleted. Taken together, these results hint at the implication of pancreatic RAS activation by ACE/Ang-II axis as a possible mechanism in diabetes development and propose ACE2/Ang-(1-7) axis as therapeutic target to help to reduce diabetes incidence, as it has been recently reviewed by Graus-Nunes and Souza-Mello [43].

Within the kidney, podocytes express ACE2 which constitutes a key enzyme within the regulation of the RAS and diabetes [44, 45]. In NOD mice, we previously observed a decrease in podocyte number at 40 days (late) of diabetes without any differences at 21 days (early) [17]. The current study demonstrated that ACE2 deletion accelerates renal disease progression in NOD mice in terms of decreased podocyte number early-on, at 30 days of diabetes. Regarding renal function, GFR in NOD.ACE2/−, tend to decrease as compared with NOD.ACE2+/+. To try to link this decrease with a faster progression of the DN we should have collected GFR measurements at 15 days of follow-up, and this approach would have helped us to detect a possible hyperfiltration state before the decrease. In the same line, UAE should be detected in early time points of the study to look for differences in filtration. Conversely, human ACE2 overexpression in podocytes leads to attenuated DN development in STZ-diabetic mice [26]. In this study, investigators showed that the improvement can be explained by a delay on albuminuria and podocyte loss prevention. The observed results suggest that the amplification of ACE2 may be a therapeutic strategy against DN and it may be indicated in early DN. Gene therapy aimed to increase ACE2 in experimental DN have been tested, however to our knowledge no beneficial effects have been demonstrated [46] since therapy failed to deliver ACE2 to the kidney.

ACE2 deletion modified RAS within the kidney. Circulating ACE enzymatic activity was increased in diabetic group as compared with controls both in ACE2KO and WT mice. In concordance, previously studies have demonstrated an increase in circulating ACE activity in STZ-diabetic hypertensive mRen2.Lewis rats and in human type 1 DM [47, 48]. In the NOD.ACE2−/− mouse line, a significant decrease in the circulating ACE activity as compared with WT diabetic mice was observed. These results indicate that the expected Ang-II accumulation in this model might activate negative feedback mechanisms to maintain homeostasis.

For the present study we isolated renal proximal tubular cells from the new NOD.ACE2−/− mouse model that we created and their respective WT mice. Our in vitro studies with renal PT cells confirmed our previous in vivo results, and demonstrated that ACE2 deletion imbalance RAS gene expression in NOD mice in terms of increasing angiotensinogen and ACE, and decreasing AT2R. This experiment confirms the effect of ACE2 modulation on RAS and consequently on the Ang-(1-7)/Mas axis. Furthermore, the strong increase in collagen gene expression under ACE2 deletion without the influence of high glucose levels is in concordance with previous studies from Oudit et al. where they demonstrated that collagen I was increased in kidneys from ACE2KO nondiabetic mice [49].

In summary, the new mouse model of spontaneous type 1 DM carrying a deletion in Ace2 gene and its nondiabetic control were viable and fertile. ACE2 loss in the NOD model leads to an impaired glucose and insulin homeostasis. This deleterious effect was accompanied by RAS activation, increase in oxidative stress, and necroptosis mediator within pancreas. In addition, any clinical situation where ACE2 is decreased may induce podocyte loss, RAS modulation, and renal fibrosis activation in an early phase of the disease.