β-Cell Deficit Due to Increased Apoptosis in the Human Islet Amyloid Polypeptide Transgenic (HIP) Rat Recapitulates the Metabolic Defects Present in Type 2 Diabetes

The generation of the HIP rats has been previously described in detail (28). Rats were bred and housed at the University of California Los Angeles animal housing facility. All animals were housed in pairs (aged 2–5 months) or in individual cages (aged 5–10 months), fed Rodent Diet 8604 (50% carbohydrate, 24% protein, and 4% fat; Harlan Teklad, Madison, WI) ad libitum and subjected to the standard 12-h light-dark cycle. The University of California Los Angeles Institutional Animal Care and Use Committee approved all surgical and experimental procedures. One week before the study, rats were anesthetized (3:3:1 Ketamine HCl, Xylazine, Acepromizine malate [0.1 ml/0.1 kg body wt, intramuscularly]) and indwelling catheters were inserted into the right internal jugular vein (dual silastic cannula, 0.03 cm [internal diameter]) and left carotid artery (polyethylene tubing, PE-50; Clay Adams 0.058 [internal diameter]). All catheters were filled with 100 units/ml heparin/saline solution, exteriorized to the back of the neck, and incased in the infusion harness. Catheters were flushed daily with 100 units/ml heparin/saline solution except on the day of clamp experiments when catheters were flushed with saline only to avoid heparin “spill over” into systemic circulation. All experimental animals were housed individually following surgical procedure.

Rats were studied 5–7 days after catheter placement in the fasting conscious unrestrained state. To assess insulin secretion, HIP (n = 17) and wild-type (n = 19) rats underwent a hyperglycemic clamp followed by an arginine bolus injection at 2, 5, and 10 months of age. On the morning of the study, animals were weighed and the venous line was connected to an adjustable infusion pump (Harvard Apparatus, Holliston, MA) for glucose infusion. The carotid line was used for blood sampling. Following a 30-min equilibration period (−30 to 0 min), animals received an intravenous glucose bolus (375 mg/kg) followed by a variable 50% (wt/vol) glucose infusion to clamp arterial glucose at ∼250 mg/dl (0–70 min). At 1 h, rats received a bolus injection of l-arginine solution (1 mmol/kg; Sigma, St. Louis, MO). Arterial blood samples (50 ul) were taken at baseline and every 10 min during the clamp for immediate determination of plasma glucose. Additional blood samples (100 ul) were collected in prechilled heparinized microcentrifuge tubes containing protease inhibitor cocktail solution (Sigma) at −30, 0, 5, 15, 30, 45, 60, 61, and 70 min and immediately centrifuged. Plasma was immediately frozen on dry ice and stored at −80°C for determination of plasma insulin, glucagon, and IAPP concentrations.

Using the same experimental setup, a different group of HIP (n = 21) and wild-type (n = 19) rats (aged 2, 5, and 10 months) underwent a hyperinsulinemic-euglycemic clamp to examine insulin sensitivity. Following an equilibration period, a constant infusion of regular human insulin (4 mU · kg⁻¹ · min⁻¹ Novolin; Novo Nordisk, Princeton, NJ) was initiated and continued until the end of the clamp (0–120 min). Plasma glucose levels were determined every 10 min, and glucose (50% wt/vol) was infused (5–120 min) to clamp plasma glucose levels at ∼100 mg/dl. Additional blood samples (∼100 μl) were collected at baseline (−30 min) and during the clamp (10, 30, 50, 80, and 120 min) for determination of plasma insulin and free fatty acid concentrations. Rates of exogenous glucose infusion were recorded, and whole-body insulin sensitivity was assessed. At 10 months of age in four of the eight HIP rats, blood glucose did not decrease during the insulin infusion at 4 mU · kg⁻¹ · min⁻¹ (i.e., they had severe hepatic and extrahepatic insulin resistance, and the glucose infusion rate would have been equal to zero). We did not include these rats in the clamp data in Table 2, which therefore report a conservative estimate of the extent of insulin resistance at 10 months. Once it was established that HIP rats develop insulin resistance by 5 months of age, basal and insulin-stimulated glucose turnover in six additional 5-month-old HIP versus five control rats were quantified using a [3-³H]glucose infusion along with the same hyperinsulinemic-euglycemic clamp protocol. Briefly, rats received primed (3 μCi)- continuous (0.05 μCi/min) infusion of [3-³H]glucose (Perkin Elmer, Boston, MA) for 90 min of an equilibration basal period followed by 120 min of tracer infusion (0.2 μCi/min) throughout the hyperinsulinemic-euglycemic clamp. Blood samples (∼150 μl) for determination of tracer specific activity at basal were drawn at − 40, −30, −20, and −10 min and during insulin infusion at 90, 100, 110, and 120 min. All blood samples were immediately centrifuged and plasma stored at −80°C for subsequent analysis.

Plasma glucose concentrations were measured by the glucose oxidase method (Beckman Glucose Analyzer 2; Beckman Coulter, Fullerton, CA). Plasma insulin, IAPP, and glucagon levels at baseline and during the hyperglycemic clamp were measured using LINCOplex multiplex rat endocrine immunoassay panel (Linco Research, St. Charles, IL). Plasma insulin concentrations during the hyperinsulinemic-euglycemic clamp were measured using competitive colomiteric enzyme-linked immunosorbent assay (Linco Research). Plasma free fatty acid levels were measured using in vitro standard enzymatic colomiteric method (Wako Chemicals, Richmond, VA). To determine plasma glucose specific activity, plasma samples were deprotonized in duplicates with ZnSO₄ and Ba(OH)₂, evaporated to remove ³H₂O, reconstituted with distilled H₂O, and assayed for ³H activity (disintegrations per mintue [dpm] per minute) with a liquid scintillation counter (Packard Bioscience, Boston, MA) using Ecoscint-H liquid scintillation fluid (National Diagnostics, Atlanta, GA). Glucose specific activity reached steady state in both HIP rats and wild-type controls in the basal and clamp sampling periods (final 30 min of baseline and clamp periods). Hepatic glucose production at baseline was assumed to equal basal R_a and, during the hyperinsulinemic-euglycemic clamp, was determined by subtracting average exogenous glucose infusion rates during the last 30 min of the clamp from the isotopically measured R_a.

After the rats were killed, pancreata were rapidly removed and all nonpancreatic tissue was also removed. Then, each pancreas was weighed and immediately fixed in 4% paraformaldehyde overnight at 4°C. The pancreas was embedded in paraffin and, subsequently, complete longitudinal sections (4 μm) of pancreas (head, body, and tail) were obtained through its maximal width. These sections were stained for hematoxylin/eosin, insulin (guinea pig anti-insulin, 1:100; Zymed, Carlsbad, CA), glucagon (rabbit antiglucagon, 1:500; Immunostar, Hudson, WI), and Congo red (Sigma), as previously described (26). The β- and α-cell mass were measured by first quantifying pancreatic cross-sectional area positive for insulin or glucagon and multiplying this by pancreatic weight. In addition, to quantify β and α apoptosis, sections were costained by immunofluorescence for insulin (guinea pig anti-insulin, 1:100; Zymed) or glucagon (rabbit anti-glucagon, 1:500; Immunostar) along with terminal deoxynucleotidyl transferase biotin-dUTP nick-end labeling (TUNEL) method for apoptosis using the cell death detection kit TMR Red from Roche Diagnostics (Mannheim, Germany) and nuclear marker 4,6-diamidino-2-phenylindole. All islets per pancreatic section (three sections per animal) were examined in detail at ×200 magnification (×20 objective, ×10 ocular) for the total number TUNEL-positive β- or α-cells, which was expressed in relation to the total number of β- or α-cells. These analyses were performed in ∼50 islets per rat (all islets on two slides per rat).

Statistical analysis was performed using ANOVA analysis and regression analysis using Statistica (version 6; Statsoft, Tulsa, OK). Data in graphs and tables are presented as means ± SE. Findings were assumed statistically significant at the P < 0.05.

There was a progressive increase in glucose concentrations in HIP rats from 2 to 10 months of age (P < 0.05; Fig. 1). At 2 months, glucose concentrations in HIP and wild-type rats were comparable; by 5 months, HIP rats had developed impaired fasting plasma glucose (128.3 ± 5.7 vs. 102.8 ± 2 mg/dl; P < 0.05); and by 10 months, overt diabetes had developed (185.3 ± 25 vs. 100.4 ± 1.5 mg/dl; P < 0.05). Fasting insulin concentrations were comparable in HIP and wild-type rats at 2 and 5 months of age, although, at 5 months, this was in the context of higher glucose concentrations (Fig. 1). By 10 months of age, insulin concentrations were deficient in HIP rats (182 ± 37.3 vs. 379.9 ± 45.6 pmol/l; P < 0.05) despite higher glucose concentrations. Fasting glucagon concentrations were comparable in HIP and control rats at 2 and 5 months. HIP rats developed hyperglucagonemia coincident with onset of diabetes at 10 months (244.4 ± 37 vs.145.3 ± 23.8 ng/l; P < 0.05; Fig. 1). Fasting plasma IAPP levels were not significantly different between HIP and wild-type rats at any point from 2 to 10 months of age (P > 0.05 for all age-groups).

Pancreas weight was not significantly different between HIP rats and controls at any age. β-Cell mass was comparable in HIP and wild-type rats at 2 months but subsequently declined progressively in HIP rats to ∼50% of control at 5 months and to ∼80% of control by 10 months of age (Fig. 2). As expected, the decline in β-cell mass in HIP rats was due to increased β-cell apoptosis, whereas α-cell mass and α-cell apoptosis were comparable in HIP rats and wild-type rats at all ages (Fig. 2).

By design during the clamp, plasma glucose levels were maintained from 5 to 70 min at ∼250 mg/dl, and clamp glucose levels were not significantly different between HIP and wild-type rats at any time point during hyperglycemia (Fig. 3). The mean glucose infusion rates required to achieve and maintain hyperglycemia were 40, 44, and 76% lower in HIP versus wild-type rats at 2, 5, and 10 months of age (P < 0.05 for all age-groups). Despite similar glucose concentrations, first- and second-phase insulin secretion to hyperglycemia was deficient and progressively declined in 2-, 5-, and 10-month-old HIP rats (∼50, 70, and 80% deficit versus wild type, respectively; P < 0.05 for all age-groups) (Fig. 3 and Table 1). Insulin secretion to arginine was not significantly different in HIP rats at 2 months of age but thereafter demonstrated a progressive decline at 5 months (1,049.1 ± 265 vs. 2,421 ± 457 pmol/l; P < 0.05) and 10 months (740.6 ± 223.4 vs. 2,478.2 ± 432 pmol/l; P < 0.05) of age.

These studies were undertaken to evaluate whole-body insulin sensitivity in HIP and wild-type rats from 2 to 10 months of age. Table 2 provides corresponding mean plasma glucose, insulin, and free fatty acid concentrations during the hyperinsulinemic-euglycemic clamp. Plasma insulin concentrations were comparable during the insulin infusions in HIP and wild-type rats at all age-groups. Whole-body insulin sensitivity, assessed by the mean glucose infusion rates during the hyperinsulinemic period, was comparable in HIP and control rats at 2 months of age (93.8 ± 7.1 vs. 100.1 ± 8.8 μmol · kg⁻¹ · min⁻¹; P = NS). However, by 5 months of age, HIP rats had an ∼40% deficit in insulin sensitivity (Table 2 and Fig. 4; P < 0.05). By 10 months of age, insulin sensitivity was markedly decreased such that the insulin infusion of 4 mU · kg⁻¹ · min⁻¹ had no effect on the blood glucose at ∼250 mg/dl in four of the eight HIP rats. These rats are not included in the mean glucose infusion rates in Table 2, so the reported glucose infusion rate is a conservative estimate of the insulin resistance in the HIP rat by 10 months of age, since only the most insulin-sensitive half of rats was included. Despite this, insulin sensitivity was still further diminished by 10 months of age in HIP versus wild-type rats (46.8 ± 3 vs. 26.46 ± 7.4 μmol · kg⁻¹ · min⁻¹; P < 0.05). When further clamp studies were performed with the isotope dilution technique in 5-month-old HIP versus wild-type rats to establish the site (hepatic versus extrahepatic) of this insulin resistance, as expected in control rats, hepatic glucose release was substantially suppressed by increased insulin during the hyperinsulinemic clamp (27.5 ± 3.25 vs. 7.52 ± 4.2 μmol · kg⁻¹ · min⁻¹; P < 0.001, basal vs. clamp; Fig. 5). However, there was a marked defect in suppression of hepatic glucose release at the same insulin concentrations in the HIP rat (34.45 ± 2.7 vs. 27.90 ± 4.5 μmol · kg⁻¹ · min⁻¹; P > 0.05, basal vs. clamp; Fig. 5), indicating hepatic insulin resistance. In contrast, at the same age, isotopically measured extrahepatic glucose uptake was comparable in HIP and wild-type rats (83.51 ± 8.5 vs. 87.48 ± 10 μmol · kg⁻¹ · min⁻¹; P > 0.05; Fig. 5), implying that extrahepatic insulin sensitivity is normal in the HIP rat at 5 months of age. Since the same insulin infusion (4 mU · kg⁻¹ · min⁻¹) had no effect on blood glucose at 10 months of age in four of the HIP rats and a much reduced effect in the four HIP rats with the least insulin resistance, by 10 months, they clearly had marked global (hepatic and extrahepatic) insulin resistance.

We report here that HIP rats and humans with type 2 diabetes not only share close islet morphology but also develop a very comparable profile of metabolic defects. In the HIP rat (and in first-degree relatives of type 2 diabetes), defective glucose-induced insulin secretion precedes hyperglycemia (8,9). In both humans and the HIP rat with IFG, there is an ∼50% deficit of β-cell mass and hepatic insulin resistance (12,29). In both humans and the HIP rat, type 2 diabetes is characterized by an ∼70% loss of β-cell mass due to increased β-cell apoptosis with a further decline in β-cell function and insulin sensitivity and hyperglucagonemia (7,12,29,30). The HIP rat is therefore an excellent model in which to explore the relationship between development of the islet pathology and metabolic changes characteristic of type 2 diabetes.

In both humans at risk of developing type 2 diabetes (8,9) and in the HIP rat, defective glucose-induced insulin secretion precedes development of type 2 diabetes. Defective glucose-induced insulin secretion was present in the HIP rat when β-cell mass was still comparable with that in wild-type rats, although the frequency of β-cell apoptosis was already increased. In the HIP rat, the increased frequency of β-cell apoptosis is related to aggregation of human IAPP (28), which has been shown to have its adverse action through toxic oligomers (18). The underlying basis of the defective glucose-induced insulin secretion in type 2 diabetes is complex (31). Once diabetes develops, hyperglycemia, per se, likely contributes to both loss of β-cell mass and function (32) in type 2 diabetes. Factors that have been implicated as mediators of this include oxygen free radical toxicity (33), expression of interleukin-1β (34), depletion of insulin stores (32), decreased flux of glucose through glucokinase (35), and mitochondrial dysfunction (36). β-Cell cytotoxic effects of free fatty acids have also been implicated, particularly in obese rodent models of diabetes, such as the diabetes-prone Zucker fatty rat (13) and the leptin-insensitive (37) or -deficient (14) mouse models. While these rodents also have a progressive defect in β-cell mass, they generally require a markedly obese phenotype before developing diabetes, and the islet does not develop the islet pathology (islet amyloid) characteristically present in most humans with type 2 diabetes. Also, the marked insulin resistance required to induce a β-cell defect in these models confounds studies of the evolving metabolic consequences of a deficit in β-cell mass.

The HIP rat has the advantage of developing diabetes with an islet pathology that is comparable with that in humans with type 2 diabetes and does not require extreme obesity. Furthermore, accumulation of misfolded proteins may provoke endoplasmic reticulum stress–induced apoptosis (38). Since protein synthesis of major client proteins by secretory cells is adaptively decreased in response to endoplasmic reticulum stress (39), this may provide a possible common explanation for the early deficit in glucose-mediated insulin secretion in the HIP rat and humans at risk of developing type 2 diabetes. Soluble IAPP may inhibit insulin secretion in a paracrine manner (40,41), and so increased expression of IAPP per cell in humans and in HIP rats could also contribute to impaired glucose-induced insulin secretion. In the HIP rat, while glucose-mediated insulin secretion was abnormal when β-cell was still unchanged (2 months), defective arginine-induced insulin secretion was not present until β-cell mass was defective (5 months), consistent with prior studies in pigs (42) reporting that arginine-induced insulin secretion more closely relates to β-cell mass than glucose-induced insulin secretion.

There is increasing interest in the pathophysiology of IFG and the transition from IFG to type 2 diabetes, since it is now possible to delay and/or prevent the development of type 2 diabetes in patients with IFG and/or glucose intolerance (43,44). In an effort to explain type 2 diabetes development, one model (45) proposed that insulin resistance eventually leads to β-cell failure and hyperglycemia. This model gains support from rodents characterized by genetic obesity that develop diabetes (13,14,37) and the fact that insulin resistance is a risk factor for type 2 diabetes (3). The problem with this model is that the majority of individuals with insulin resistance, most commonly due to obesity (1,2), adaptively increase insulin secretion (6,7) and do not develop diabetes (5). Therefore, the role of insulin resistance in the pathogenesis of type 2 diabetes appears to be to induce the β-cell defect in those who are genetically vulnerable. One mechanism by which this might theoretically be accomplished is the increased expression of IAPP that occurs in response to insulin resistance (46). Previously, we proposed that there might be a lower capacity to fold and traffic IAPP in genetically vulnerable individuals (e.g., as a result of polymorphisms in chaperone proteins); this concept is gaining acceptance in other similar amyloidogenic diseases such as Alzheimer’s disease (47,48). The HIP rat provides a powerful tool to dissect out the importance of IAPP expression per se versus insulin resistance on progressive islet dysfunction. Thus, at 2 months of age, the HIP rat has increased human IAPP expression because of human IAPP transgenic expression rather than insulin resistance. Indeed, at 2 months of age, both blood glucose and insulin sensitivity are normal in the HIP rat, but by 5 months of age, the HIP rat develops hepatic insulin resistance coincident with the onset of IFG, and by 10 months of age, with further progression of insulin resistance, the HIP rat develops diabetes. These data imply that hepatic insulin resistance characteristic of IFG (29) and hepatic and extrahepatic characteristic of type 2 diabetes (30) might at least in part be due to β-cell failure. In the context of the population that develops type 2 diabetes most frequently already being insulin resistant (obese), we propose that the additional insulin resistance in individuals with IFG and type 2 diabetes versus comparably obese nondiabetic control subjects may be at least in part due to β-cell failure (29,30).

In healthy subjects, insulin is secreted in discrete insulin secretory bursts that present the liver with an insulin concentration profile that oscillates with an amplitude of ∼1,000 pmol/l in the fasting state (49,50) and up to 5,000 pmol/l after meal ingestion (51). As β-cell mass is decreased in animal models, both basal and glucose-stimulated insulin secretion decline due to a selective defect in insulin pulse mass (52,53), a profile that has been observed in humans with type 2 diabetes (54). The pulsatile mode of insulin delivery to the liver increases hepatic insulin sensitivity (55,56), possibly by avoiding insulin receptor downregulation (57) and/or sustained activation of AKT with attendant feedback inhibition of insulin receptor substrate-2 (58). We postulate that the decline in hepatic insulin sensitivity in the HIP rat (and in humans) with IFG may relate at least in part to the decline in β-cell mass leading to an abnormal pattern of insulin delivery to the liver. Since hepatic insulin extraction depends on the amplitude of insulin pulses presented to the liver (59), this defect in insulin secretion would be largely offset in the systemic circulation by decreased hepatic insulin clearance. Another theoretical explanation for insulin resistance in the HIP rat is the actions of circulating IAPP, since high IAPP concentrations have been shown to cause insulin resistance (60). However, this effect has been shown in muscle rather than liver (61) and requires IAPP concentrations in the nanomolar versus low picomolar range observed here (Fig. 1). Moreover, the circulating IAPP concentrations were not increased in HIP versus wild-type rats (P > 0.05 for 2–10 months of age). As expected from prior studies (62), once diabetes supervened at 10 months of age, extrahepatic insulin resistance was also present, presumably as a consequence of the effects of glucose toxicity (63). Indeed, once HIP rats developed overt hyperglycemia (>250 mg/dl) from 12 to 18 months of age, the insulin infusion rate (4 mU · kg⁻¹ · min⁻¹) used during the hyperinsulinemic-euglycemic clamp was no longer sufficient to lower plasma glucose in these animals to ∼100 mg/dl.

There are limited data regarding the importance of β-cell mass in the pathophysiology of IFG or type 2 diabetes because of the paucity of data on β-cell mass in humans. These data are only available from autopsy studies, revealing an ∼50 and 70% deficit in β-cell mass in IFG and type 2 diabetes, respectively (12), but, by necessity, these data are only available at a single time point. When the relationship between blood glucose and β-cell mass in humans is examined, a β-cell mass of ∼50% of normal level is the transition between hyperglycemia development vulnerability and a capacity to secrete sufficient insulin to maintain normal blood glucose concentrations (64). It is therefore of interest that the curve describing the relationship between β-cell mass and fasting blood glucose in the wild-type and HIP rat also shows a point of inflection at ∼50% of the mean of β-cell mass in wild-type rats (Fig. 6). As in the human curve, there is a steep increase in blood glucose with each decrement of β-cell mass thereafter, consistent with the known adverse effects of hyperglycemia on β-cell function and survival (34). There are comparable cross-sectional data in dogs (65), monkeys (66), and pigs (53) showing that an ∼50% deficit in β-cell mass is sufficient to lead to IFG. Also, living related donors of individuals with type 1 diabetes who had a 50% pancreatectomy had a high risk of subsequently developing diabetes (67). The problem with prior data is that they are cross-sectional, so it is not possible to be certain that humans developing type 2 diabetes progress from a normal β-cell mass through to ∼50% deficit with IFG and ∼70% deficit with diabetes. While it is not possible to be certain that the same sequence of defects in β-cell mass and metabolic dysfunction occur in humans with type 2 diabetes and HIP rats, before we can quantify β-cell mass in humans, the HIP rat allows us to examine the evolution of an islet pathology closely resembling that in humans with IFG and type 2 diabetes. Simultaneously, the HIP rat allows us to quantify insulin secretion and insulin action and imply that it is certainly possible that the β-cell deficit in humans with type 2 diabetes is both progressive and closely related to the development of the disease.

In the present study, we also report that in contrast to the β-cell mass, the α-cell mass in HIP rats does not differ from that in wild-type rats from 2 through 10 months of age (i.e., throughout the transition from the nondiabetic state to IFG and diabetes). Consistent with this finding, the measured frequency of α-cell apoptosis did not change throughout this period. These data are consistent with the selective deficit in β-cell mass reported in humans with type 2 diabetes (12). They also emphasize that IAPP-induced β-cell death and the associated islet amyloid does not influence viability of other islet cells, reinforcing the accumulating evidence that it is not extracellular amyloid that is cytotoxic but rather intracellular toxic oligomers of amyloidogenic proteins (18). An alternative explanation is that α-cells are resistant to amyloid-related toxicity, although this is unlikely since the addition of amyloidogenic proteins to many different cell types in culture has been reported to induce apoptosis (68–70). We also report here that in the HIP rat, hyperglucagonemia develops coincident with the development of diabetes. From these data alone, it is not possible to determine if this is due to declining intra-islet insulin-mediated inhibition of glucagon secretion or secondary to hyperglycemia per se. Nonetheless, this finding again provides a parallel to humans with type 2 diabetes who have hyperglucagonemia that contributes to postprandial hyperglycemia (29).

In summary, in the HIP rat, we report that the β-cell human IAPP transgene leads first to increased β-cell apoptosis and β-cell dysfunction (age 2 months), then to hepatic insulin resistance leading to IFG (5 months), and finally to more severe insulin resistance, hyperglucagonemia, and diabetes (10 months). Through this transition, we document both a progressive decline in glucose-mediated insulin secretion (both first and second phase) and a relative decline in β-cell mass that is comparable in extent to that in humans with IFG (∼50%) and type 2 diabetes (∼70%), respectively (12). Of particular note, this selective deficit in β-cell function, and then mass, leads to hepatic insulin resistance. These data imply that the added insulin resistance characteristic of type 2 diabetes may be a consequence, rather than the cause, of β-cell dysfunction.

We gratefully acknowledge funding for these studies from the U.S. Public Health Service (National Institutes of Health DK59579) and the Larry Hillblom Foundation.

We gratefully acknowledge the helpful suggestions from our collaborators at the Larry Hillblom Islet Research Center, UCLA Anil Bhushan, Alexandra Butler, Tatyana Gurlo, Leena Haataja, Kathrin Maedler, and Walter Soeller at Pfizer Research in Groton, Connecticut.