Background

The cysteine proteases calpain and caspase-3 are known mediators of cell death. The aim of this study was to assess their contribution to the tissue damage found in experimental uremia.

Methods

Calpain and caspase-3 activities were measured in the hearts of rats that were sham-operated (control), sham-operated and spontaneously hypertensive (SHR), and those rendered uremic by 5/6 nephrectomy (uremic). In an in vitro study, heart myoblasts (Girardi) were incubated with human serum from healthy subjects (control serum conditioned media, CSCM) or uremic patients (uremic serum conditioned media, USCM), in the presence and absence of calpain and caspase-3 inhibitors. After 48 hours the activity of calpain and caspase-3 was measured, and cell injury determined by DNA fragmentation (ELISA) and lactate dehydrogenase (LDH) release. An in situ assay was designed to study how USCM affects calpain activity over time.

Results

In the in vivo study, mean calpain activities were almost identical in the control and SHR groups, but calpain and caspase-3 activities were much elevated in the uremic group (P < 0.01 and 0.001 respectively vs. control). The SHR group had significantly higher mean arterial blood pressure (P < 0.001 vs. control, 0.01 vs. uremic). In the in vitro study calpain activity and DNA fragmentation were markedly higher in USCM treated cells compared to CSCM (both P<0.05). Both were reduced in USCM cells containing calpain inhibitors (E64d, calpastatin, or PD 150606). LDH release was raised also in USCM treated cultures (P < 0.05), which only the E64d treatment could significantly reduce (P < 0.02). Caspase-3 activities were similar in USCM and CSCM groups. The in situ assay showed significant increases in calpain activity in USCM treated cells compared to CSCM after just 3.5 hours (P<0.01).

Conclusions

In vivo results suggest that the increases in calpain and caspase-3 activity in uremic rat hearts were primarily due to uremia and not to hypertension. In vitro data demonstrate that uremia-induced cell injury can be attenuated by calpain inhibition. Therefore, it is likely that calpain is a mediator of uremia-induced myocardial injury.

METHODS

Animals

All animal procedures were ethically approved and performed under current United Kingdom Home Office regulations (Project license number 70/5014). Male Wistar Kyoto (WKY) rats and the genetically related SHR strain were obtained from Harlan UK Ltd. (Bicester, Oxon, UK). One group of rats was made uremic by 5/6 nephrectomy using an established two-stage surgical procedure (uremic)¹⁴. A second group of WKYs were sham-operated upon by removing the renal capsule and replacing the intact kidney (controls), while a third group consisted of sham-operated SHRs (N = 7 for all three groups). SHR and control animals were pair-fed with the sub-nephrectomized animals to ensure that significant differences in body weights did not occur. The study lasted for 12 weeks post-surgery. Rats were housed individually and given water ad libitum. Mean arterial blood pressure (MAP) was determined on conscious unrestrained animals intra-arterially (see below). At the end of the study the animals were killed by cervical dislocation, and the hearts were promptly removed and kept frozen at -80°C.

Measurement of mean arterial pressure

Femoral artery cannula insertion was performed 24 hours prior to blood pressure recordings. Measurements were made under quiet resting conditions with the rat free to move within the cage. To record blood pressure the catheter was clamped while the distal (knotted) end was cut off with scissors, then the catheter was connected to a pressure transducer (Model number 60-3003; Harvard Apparatus, Boston, MA, USA) through a length of tubing, and the clamp released. An analogue-to-digital converter (Dataq Instruments Inc., Akron, OH, USA) allowed the output from the pressure transducer to be continuously displayed and recorded on a PC using Windaq (Dataq Instruments Inc.) digital chart recorder software. MAP was determined by placing a 1 mL air-filled tube in parallel with the arterial line through a three-way tap; arterial pressure traces were adequately damped to give a virtually flat mean arterial signal. The arterial cannula was then used to acquire blood for analysis.

Cell culture

Human-derived Girardi cells were obtained from the European Collection of Animal Cell Cultures (ECACC, Porton Down, UK). Cells were cultured in EMEM media containing 2 mmol/L glutamine, 10% fetal calf serum (FCS), 1% non-essential amino acids (NEAA) from GIBCO™ (Life Technology Ltd, Paisley, UK) and antibiotic (100 U/mL penicillin, 100 g/mL streptomycin and 0.25 g/mL amphotericin B) in 25 cm³ tissue culture flasks. Cultures were maintained in media containing 10% FCS while experiments were conducted in media containing 5% FCS and 5% human serum from healthy volunteers or patients with renal failure.

In vitro model of uremia

Serum was obtained from patients with ethical approval and informed consent (ELCHA Research Ethics Committee, P/97/334). All patients were treated by hemodialysis, not on antihypertensive therapy, and samples were taken just prior to dialysis. Control samples were from normotensive individuals and matched for gender (all male), but not for age (in years SE, controls = 38.8 2.3, uremic patients = 50.1 6.2, mean time on dialysis 2.8 0.6). However, the ages of the two groups were not statistically different (P = 0.086, N = 8). Serum samples from each individual was prepared in media (0.5 mL FCS, 0.5 mL human serum, 9 mL of EMEM media) and was divided into two flasks of subconfluent cells (cells confluent at 48 hours), one of which was used as a time 0 hour baseline.

Calpain inhibitors and the caspase-3 cell permeable inhibitor DEVD-CHO (CN Biosciences UK Ltd, Nottinghamshire, UK) were added to uremic serum conditioned media (USCM) at the start of the experiment. The concentration of inhibitors was determined from the highest effective concentration that did not produce cell injury in the cells treated with normal serum conditioned media (CSCM). This was because, although high concentrations of PD 150606 (100 mol/L) could completely inhibit calpain activity in vitro, it also caused a marked increase in lactate dehydrogenase (LDH) release (data not shown). After (0 and) 48 hours of incubation at 37°C in a humidified environment of air/CO₂ (19:1), cells were scraped off and centrifuged (100 g, room temperature for 5 min). An aliquot of each CSCM and USCM used was reserved prior to the experiment, at times 0 and 48 hours, and then stored at -20°C for the LDH assay. Cell pellets were resuspended in 400 L of cold assay buffer (63.2 mmol/L imadazole-HCl containing 10 mmol/L of 2-mercaptoethanol, pH 7.3) containing 10 mol/L digitonin, 1 mmol/L egtazic acid (EGTA), 0.1 mmol/L ethylenediaminetetraacetic acid (EDTA; pH 7.3), and kept on ice for 30 minutes with frequent mixing. Samples were then centrifuged (14,000 g, 4°C, 30 min; Sorval RMC 14), the cell pellets discarded, with the lysates kept on ice until assayed for calpain, before being frozen at -20°C.

Calpain activity assays

The assays described in our study used synthetic peptides as substrates that contain a molecule that fluoresces only after protease cleavage. The peptides were not specific for calpain, so the assay conditions were manipulated to ensure that only the activity of calpain was measured. This involved the use of physiological conditions (where cathespins were known to be inactive), the measurement of calcium dependent activity only (cathespins and caspases were calcium-independent), or as in the case of the in situ assay (calpain activity in situ) by the use of calpain-specific inhibitors. The use of these peptides for the measurement of calpain has recently been reviewed¹⁵. With the exception of the in situ assay, calpain activity was determined as the difference between the calcium dependent fluorescence and the non-calcium dependent fluorescence. A 7-amino-4-methylcoumarin (AMC; ICN Biomedicals Ltd., Thame, Oxfordshire, UK) standard curve was constructed for each assay with each calibrant containing the same concentration of DMSO as the samples. Calpain activity was expressed as nmol of AMC released per minute of incubation time per mg of total protein.

Calpain activity in rat hearts

Calpain activity was determined using an assay procedure that was modified from those described by Sasaki et al for measurement in purified porcine kidney¹⁶ and by Edelstein et al in rat proximal tubules¹⁷. Briefly, the assay was performed as follows: Assay buffer consisted of 63.2 mmol/L imadazole-HCl containing 10 mmol/L of 2-mercaptoethanol (pH 7.3). Calcium-free buffer was prepared in assay buffer containing 20 mmol/L EGTA and 25 mmol/L EDTA (pH 7.3). Calcium buffer consisted of assay buffer with 2.5 mmol/L CaCl₂ added (pH 7.3). Freshly thawed hearts were weighed prior to the addition of chilled calcium-free buffer (5 mL/g tissue). Tissue was disrupted with a sintered glass homogenizer prior to centrifugation (14,000 g, 4°C, 30 min; Sorval RMC 14). The assay was then performed on this supernatant after it was diluted fivefold in calcium-free buffer. To four tubes containing 500 L of diluted supernatant, one pair had 1.5 mL of calcium-free buffer added, and the other pair had the same volume of calcium buffer added. After a 10 minute pre-incubation period of shaking in a water bath at 37°C, 10 L of the substrate N-succinyl-Leu-Tyr-AMC (10 mmol/L in DMSO) was added to all tubes. After 30 minutes of incubation fluorescence was detected at 380 nm excitation and 460 nm emission (Shimadzu RF-5000; Shimadzu UK Branch, Milton Keynes, UK).

Calpain activity in cell lysates

The assay described in the last section was further modified in order to determine calpain activity in cell lysates. Lysates were kept on ice during assay. Calcium-free buffer was prepared in assay buffer containing 10 mmol/L EGTA and 1 mmol/L EDTA (pH 7.3). Calcium buffer consisted of assay buffer with 5.0 mmol/L CaCl₂ added (pH 7.3). Sample lysate (10 L) was added to four wells of a black polypropylene microtiter assay plate, followed by 90 L of calcium-free buffer to two of the wells and 90 L of calcium buffer to the remaining pair of wells. After 10 minutes of pre-incubation with shaking in an incubator at 37°C, 2 L of the substrate N-succinyl-Leu-Leu-Val-Tyr-AMC (1 mmol/L in DMSO) was added to all wells. Fluorescence was detected at 380 nm excitation and 460 nm emission on a microtiter fluorescence plate reader (BMG Fluorostar; BMG Labtechnologies Ltd, Aylesbury, UK).

Calpain activity in situ

A 25 cm³ tissue culture flask of sub-confluent cells were treated with 5 mL of 0.25% trypsin EDTA and seeded into a 96 well tissue culture plate with 150 L of EMEM containing 10% FCS media/well. After 24 hours the wells consisted of a monolayer of cells. The cell culture media was then removed and replaced with 150 L of human serum enriched media solutions (N = 7 in each group) containing 5% FCS supplemented with 5% of either control (CSCM) or uremic serum (USCM). Each media solution was added to 8 wells of the tissue culture plate containing Girardi cell monolayers and a blank plate (without cells). To both plates 50 mol/L of the calpain-specific inhibitor calpeptin (in DMSO) was added to four wells, while the remaining 4 wells were treated with the same quantity of DMSO (2 L, 1.33%). After 10 minutes of pre-incubation at 37°C, 2 L of the substrate N-succinyl-Leu-Leu-Val-Tyr-AMC (1 mmol/L in DMSO) was added to all wells containing media in both plates. Fluorescence was detected at 380 nm excitation and 460 nm emission on a microtiter fluorescence plate reader. Calpain activity was determined for each media solution as the difference between the mean fluorescence in the samples without the calpain inhibitor and those with the inhibitor. The calpain activity of each media solution alone (from the blank plate) was then subtracted from the calpain activity from the tissue culture plate to ensure that only the calpain activity in the cells was measured. In addition to the human serum media samples, 8 wells of each plate were treated with media supplemented with 10% FCS (negative control), and a further 8 wells with 10% FCS containing 2 mol/L of the calcium ionophore A23187 (positive control). These samples were then treated with calpeptin (or DMSO) and substrate as described earlier.

Caspase-3 activity in tissue homogenates

The tissue homogenates were prepared as before (Calpain activity in rat hearts) and caspase-3 activity was determined by a commercially available activity assay (Promega, Southampton, Hampshire, UK). The assay was performed as stated in the kit documentation using 20 L of homogenate per tube in the standard assay format. Fluorescence was detected at 380 nm excitation and 460 nm emission on a spectrofluorophotometer (Shimadzu). Caspase-3 activity was expressed as nmol of AMC released per minute of incubation time per mg of total protein.

Caspase-3 activity in cell lysates

As described in the last section, except that 10 L of cell lysate were used, the assay protocol was then as stated in the kit documentation for the 96-well plate format. Fluorescence determined on a microtiter fluorescence plate reader.

Quantification of cell death by DNA fragmentation assay

Cell death was detected by a DNA fragmentation enzyme-linked immunosorbent assay (ELISA), which quantifies the mono- and oligonucleosomes in the cytoplasmic fraction of cell lysates or incubation media (Cell Death Detection ELISA^PLUS; Roche Diagnostics, Lewes, UK). Lysates were diluted 20-fold in kit lysate buffer while cell incubation media were sampled neat. The assay was performed as described in the kit insert with DNA fragmentation expressed as an enrichment factor (absorbance value for 48-hour experimental sample/absorbance value of 0 hour sample) from a time 0 hours baseline sample per mg of total lysate protein.

Quantification of cell death by LDH assay

Lactate dehydrogenase activity was measured in the incubation media prior to (background control) and at the end of the experiment (48 h) using a colorimetric assay (Cytotoxicity Detection Kit; Roche Diagnostics). Cytotoxicity (%) was calculated by the subtraction of the background control from the 48 hour sample and expressed as a percentage of a positive control (2% Triton X-100), minus background control, corrected for total lysate protein.

Other measurements

Total protein concentration was determined by Bradford assay and serum creatinine by the Jaffé rate method (Beckman 2 Analyzer; Beckman Coulter U.K. Ltd., High Wycombe, UK). Serum calcium and phosphate concentrations were performed by VETLAB services (Southwater, Sussex, UK).

Western blotting

Tissue homogenates were diluted tenfold in calcium-free buffer and lysates were used neat. All samples were initially diluted in running buffer (2 L buffer/8 L of sample), before being boiled at 99°C for five minutes. Samples were placed on ice before gel loading (10 g protein/well of homogenates and lysates, 20 ng of human erythrocytes calpain-1 and rat recombinant calpain-2; CN Biosciences UK Ltd, Nottinghamshire, UK). Proteins were separated on a 7.5% poly-acrylamide gel (National Diagnostics Ltd, Hessle, UK) and electro-blotted onto a polyvinylidine (PVDF) membrane (Immobion-P™, Sigma, UK) and blocked overnight at 4°C with 5% BSA. Membranes were then probed with mouse anti -fodrin monoclonal antibody diluted 1:1000 (Affiniti Research Products Ltd., Mamhead, UK) or goat anti-calpain-1 or -2 (1:1000; Santa Cruz Biotech, Autogen Bioclear, Calne, UK). The secondary antibody was HRP labeled IgG (1:2000; Santa Cruz Biotech). Comparative densitometry was performed on the fodrin 145/150 kD doublet and the calpain-2 band visualized using chemiluminescence (ECL; Autogen Bioclear).

Two of the membranes Figure 2a, c were stripped by incubation in 62.5 mmol/L Tris (pH 6.8), 2% SDS, 100 mmol/L 2-mercaptoethanol for 30 minutes at 50°C. After washing and re-blocking, the membranes were re-probed with a different primary antibody Figure 2b, d.

Figure 2.

Western blots demonstrating that control rat hearts (C heart) contain mostly calpain-2 immunoreactivity with very little calpain-1 evident (A,B). (A) Cells were stripped and re-probed with anti-calpain-2 antibody (B). Purified calpain-1 was derived from porcine erythrocytes and purified calpain-2 is rat recombinant. Both positive controls (20 ng) are indicated by dark bands when probed by the appropriate primary antibody, but only calpain-2 immunoreactivity is clearly visible in the heart homogenate. Calpain-2 activity was always much higher in uremic heart tissue, but this was not always evident from Western blots (C). (D) An increase in fodrin 145/150 breakdown products (BDPs) in uremic hearts is shown. (E) One mol/L (arrowed) of the specific calpain inhibitor PD 150606 is able to inhibit the formation of 145/150 fodrin BDPs in uremic serum conditioned myoblasts. Panels (C) and (D) portray 3 "triplets" of hearts that were representative of the 7 "triplets" studied. Panel (E) is representative of two separate experiments that were both run on two gels.

Full figure and legend (52K)

Materials

Unless otherwise stated in the text, all chemicals were obtained from the Sigma-Aldrich Chemical Company (Poole, Dorset, UK).

Statistical evaluation

All data are presented as means SE of N observations, where N represents the number of animals or serum-enriched media samples studied. Data were initially tested for normal distribution (Shapiro-Wilk test) by using Analyse-it™ software for Microsoft Excel (v1.62; Leeds, UK). Animal data were analyzed by one-way ANOVA (matched) with Tukey's post-hoc test, and cell culture data by the unpaired Student t test. When comparing the means of two groups, P < 0.05 was considered significant.

RESULTS

The 5/6 nephrectomized rats had (as expected) markedly elevated serum creatinine concentrations compared to both the control and SHR groups, thereby confirming the onset of uremia Figure 1a.

Figure 1.

Serum creatinine (A), final heart wet weight (B), conscious mean arterial blood pressure (C; MAP), homogenate calpain (D) and caspase-3 (E) activities in animals 12 weeks after the uremic animal group were 5/6^th nephrectomized (N = 7 sets of 3 animals).

Full figure (42K)

The hearts of the uremic rats were significantly larger (as determined by wet weight) than those from the control group Figure 1b while the SHR group mean heart size was not statistically greater than controls, indicating the development of uremia-induced left ventricular hypertrophy (LVH) in the subtotally nephrectomized animals.

The uremic and SHR groups both had significant higher mean arterial blood pressures (MAPs) but MAP was highest in the SHR group Figure 1c. This suggests that the SHR group can act as a control for the hypertension associated with uremia and that the effects found in this study are due predominantly to uremia and not to high blood pressure.

The hemoglobin concentrations revealed that the uremic animals were anemic (in g/dL SE, control 14.9 0.8, SHR 15.8 1.3, uremic 8.6 1.0, control vs. uremic P < 0.01, SHR vs. uremic P < 0.005, control vs. SHR = NS). There were no significant differences in the final body weights of the rats (in g SE, control 369 2.3, SHR 366 8.8, uremic 314 19.5). There were also no significant differences in serum calcium (in mmol/L SE, control 1.74 0.28, SHR 1.88 0.10, uremic 1.49 0.31) and serum phosphate concentrations (in mmol/L SE, control 3.94 1.63, SHR 2.32 0.18, uremic 4.88 1.69).

Calpain and caspase-3 activities were found to be higher in the homogenized hearts from the uremic group compared to sham-operated animals Figure 1d, and e. This was confirmed by the increase in fodrin BDP 145/150 kD doublet intensity in the uremic group compared to control animals Figures 2d and 3b. It is noticeable that the relative increase in calpain activity with respect to the control group was much greater than for caspase-3 (3.4-fold rise for calpain compared to a 2.0-fold for caspase-3). Calpain-2 immunoreactivity was clearly visible on Western blots Figures 2c and 3a and markedly more calpain-2 immunoreactivity than calpain-1 was evident in control animal hearts Figure 2a, b. The density of the calpain-2 band tended to be greater in the uremic animal group compared to control and SHR groups, but it failed to reach statistical significance (Figure 3a; N = 7). Therefore, the density of the calpain-2 band did not always reflect total calpain activity, as all of the uremic hearts had much higher calpain activities than their paired control and SHR hearts Figure 1d.

Figure 3.

Densitometry data derived from Western blots for calpain-2 bands and the fodrin 145/150 BDP doublet (N = 7). No statistical differences were found in the densities of the calpain-2 band (A). The fodrin 145/150 kDa doublet band density was significantly increased in the uremic hearts (B).

Full figure and legend (34K)

In vitro model of uremia

The mean creatinine concentration in the serum used in these cell culture experiments was 77 mol/L (5.3, SE) for the healthy volunteers, while in the patients with renal failure it was 883 mol/L (67 SE, N = 8, P < 1 10^-8).

Cell death (as determined by DNA fragmentation and LDH release) and calpain activity were significantly higher in the lysates and supernatants (incubation media) from cells treated with USCM for 48 hours compared with those treated with CSCM Figure 4a–d. The elevation in lysate DNA fragmentation and calpain activity in USCM cells were significantly reduced in USCM cells containing PD 150606, E64d Figure 4a, d, and calpastatin peptide Figure 4e, f. Calpastatin-negative control peptide (scramble peptide of identical amino acids) as predicted had no detectable effect Figure 4e, f on lysate DNA fragmentation and calpain activity. However, calpain inhibition was unable to lower LDH release in the USCM cells using PD 150606 Figure 4c or calpastatin peptide (data not shown), but E64d, a broader-based cysteine protease inhibitor, did reduce LDH release Figure 4c. Western blots confirmed the ability of PD 150606 at 1 mol/L to lower calpain activity in this model by reducing the formation of calpain/caspase-3 specific fodrin BDPs Figure 2e. Caspase-3 activity was found to be no higher in lysates from USCM treated cells and those incubated with CSCM Figure 5a, while a specific caspase-3 inhibitor had no discernable effect on the degree of DNA fragmentation Figure 5b, despite greatly reducing caspase-3 activity Figure 5c.

Figure 4.

DNA fragmentation in (A) lysates and (B) incubation media, lactate dehydrogenase (LDH) cytotoxicity in incubation media (C), and calpain activity in lysates (D) derived from Girardi cells incubated for 48 hours in healthy and uremic serum-conditioned media, with and without the calpain inhibitor PD 150606 or E64d, a cysteine protease inhibitor. DNA fragmentation and calpain activity (E and F) in uremic-serum conditioned media with and without calpastatin peptide and a negative control scramble peptide (N = 6 to 8 for all experiments).

Full figure and legend (49K)

Figure 5.

Caspase-3 activity (A) in lysates from Girardi cells incubated for 48 hours in healthy and uremic serum-conditioned media. The cell permeable caspase-3 inhibitor DEVD-CHO did not reduce the degree in DNA fragmentation induced by uremia (B) despite significantly reducing caspase-3 activity (C, N = 6 to 8).

Full figure and legend (38K)

In situ calpain assay

The in situ calpain assay revealed that calpain activity in cultured myoblasts rises sharply within hours of treatment with USCM. Figure 6a shows mean calpain activities in Girardi cells after a six hour incubation in media enriched with CSCM or USCM (N = 7), while Figure 6b shows the rise in calpain activity over time (N = 3). The addition of the calcium ionophore A23187 (1 mol/L) caused a rapid rise in calpain activity Figure 6b. A23187 is known to fluoresce, but as we detected the difference in activity in A23187 treated cells with and without calpeptin, this fact is not relevant in this system. FCS (10%) treatment promoted calpain activity in a similar manner to control media (5% FCS and 5% serum from normal subjects).

Figure 6.

In situ calpain activity in monolayers of Girardi cells incubated with media enriched with serum from healthy human volunteers (controls) or patients with end-stage renal failure (N = 7) for 6 hours (A). (B) Uremic serum-enriched media elicits a significantly higher calpain activity than control media after just 3.5 hours (210 min, P < 0.01, 250 min P < 0.02, 300 min P < 0.02, and 350 min P < 0.02; N = 3). Calpain activity rose sharply when treated with the calcium ionophore A23187.

Full figure and legend (31K)

DISCUSSION

Our study illustrates the injurious effect upon cardiac tissue of experimental uremia and that uremia causes significant increases in calpain activity. This is supported by the finding that the mean heart weight for the uremic group of animals was significantly higher than the control group Figure 1b, indicating the development of left ventricular hypertrophy (LVH) in this group of animals. LVH appears to be induced by uremia rather than high blood pressure, as the more hypertensive SHR animals had heart weights not significantly greater than control rats Figure 1b, c. The elevation of total calpain activity and caspase-3 activity in the uremic rats is unequivocal Figure 1d, e and is supported by the increase in the density of the fodrin 145/150 kD BDP (Figure 2d and Figure 3b). However, the density of the calpain-2 immunoreactive band did not always correspond with the increase in total calpain activity in the uremic heart Figures 2c, 3a, and 1d. This could possibly be due to uremia causing a decrease in endogenous calpastatin (calpain inhibitor) or an increase of other calpain isoforms. A further possibility is that the calpain present in the uremic heart is more active than that in a healthy heart. Certainly the evidence of the in situ data showing that calpain activity was significantly higher in cells incubated with USCM compared to those incubated with CSCM after just 3.5 hours Figure 6b suggests that calpain activation in uremia is not due solely to increases in calpain expression. Calpain activity was still elevated in situ after six hours Figure 6a and in the in vitro model after 48 hours Figure 4d. However, when interpreting the results of this in vivo study it is important to acknowledge the limitations of this work. Firstly, the uremic animals also were found to be anemic, and secondly, the SHR group of animals failed to develop significant LVH. Therefore, it is not possible to state that the changes in the cysteine protease activities in the uremic rat myocardium are purely as a result of uremia. This is because LVH itself and/or anemia might also play a role.

The sera used in these cell culture experiments were obtained from healthy human volunteers and uremic patients treated by hemodialysis. It is feasible that hemodialysis itself rather than uremia might contribute to the activation of calpain in vitro. Therefore, it would be of interest to examine whether serum from patients treated with continuous ambulatory peritoneal dialysis could induce calpain activation in the same manner. The mean age of the uremic patient group was higher than the control group (although not statistically), but it is doubtful that age is a factor in this study, as the two eldest uremic patients (aged 65 and 79) induced the lowest increases in calpain activity of the uremic population studied (data not shown).

The reduction of lysate DNA fragmentation in uremic serum-treated cells incubated with two distinct and specific calpain inhibitors suggests that calpain has an important role in the development of uremia-induced myocardial injury. In addition E64d (which inhibits calpain and other cysteine proteases) was able to reduce both DNA fragmentation and LDH release, thereby indicating a role for additional proteases in uremia induced cell death. The elevation of DNA fragmentation found in cell lysates Figure 4a, and incubation media Figure 4b suggests that uremia induced injury involves both apoptotic and necrotic cell death. Thus, it is possible that specific calpain inhibition can decrease uremia induced apoptosis but not necrosis. This indicates that the cells are destined to die by events upstream of calpain. This is in part analogous to the findings of Suzuki et al, who found that the inhibition of caspases in peroxide-induced apoptosis caused a decrease in apoptosis but increased the rate of necrosis¹⁸. There are many examples of calpain having a significant role in apoptosis and necrosis. In PC12 cultures, calpain has been shown to possess the more dominant role in hypoxic cell death than the caspases¹⁹, and oxidative stress induced apoptosis can be restricted by calpain inhibition²⁰. The pro-apoptotic BH3-only Bcl-2 family member protein Bid was found to be cleaved during myocardial ischemia/reperfusion to an active fragment by calpain causing cytochrome c release without caspase activation²¹. Specific calpain inhibition also has been shown to protect against virus-induced apoptotic myocardial injury in neonatal mice²².

The influence of caspase-3 in this study was harder to determine. Although the mean caspase-3 activity was significantly higher in the hearts of uremic animals compared to control rats, the fold rise was modest when compared to the increase in calpain activity. Furthermore, mean caspase-3 activities in lysates derived from cells incubated in USCM were no higher than those in cells incubated in CSCM Figure 5a. This might be because uremia is inducing cell death solely via a necrotic pathway, but the presence of DNA fragmentation in cell lysates indicates the presence of some apoptotic death. This can be likened to the findings of Ueda et al, who studied the effects of hypoxia/reoxygenation injury (which is classically described as a form of necrotic cell death) in kidney proximal tubules^23,24. They found evidence of DNA fragmentation and endonuclease activation but no morphological evidence of apoptosis. Their work concludes that when the degree of ischemia is severe the cells exhibit a necrotic form of death, and where it is mild the cells can undergo complete repair. It is when the insult is between these two extremes, and complete repair cannot be accomplished that an apoptotic cascade of events can occur. There also have been numerous reports of apoptosis that are caspase-3 independent^{25,26,27,28,29,30}. An example of this is that cadmium induced apoptosis in U937 cells appears to occur through two independent pathways, a Ca²⁺/calpain dependent pathway and a caspase-mitochondria dependent pathway²⁶. In addition Huang and Wang have suggested that ischemia and traumatic brain injuries that result in oncosis and apoptosis are mediated via calpain activation independently of caspases³. Therefore, it is possible that uremia is inducing necrotic and/or apoptotic cell death that is independent (at least in vitro) from caspase-3. It also is feasible that caspase-3 does play a role in uremia-induced cell injury in cell culture, but we did not detect it due to measuring its activity at an inappropriate time point. However, this seems unlikely, as the use of a potent cell permeable caspase-3 inhibitor was unable to reduce uremia-induced DNA fragmentation in this study Figure 5b despite reducing caspase-3 activity greatly Figure 5c. Recent work by Knaapen et al suggests that cardiomyocytes in heart failure suffer from caspase-independent autophagic cell death rather than apoptotic cell death³¹. However, the role of caspase-3 and other caspases still warrants further study because they are known to interact with calpain in a number of diverse pathways^32,33,34. Of particular relevance might be the finding that caspase-3 and procaspase-3 are substrates for calpain with calpain cleavage rendering caspase-3 (and other caspases) inactive^33,35. It might be that an elevation in caspase-3 activity in uremic serum-treated cells is being prevented by a rise in caspase-3 deactivation due to the increase in calpain activity.

The reason why calpain is activated in uremia as yet can only be speculated upon. It is possible that calcium has a role as calcium homeostasis is altered in both experimental and clinical uremia^13,14. In this study no evidence of an increase in circulating Ca²⁺ concentrations in the uremic rat was found. Future work will investigate the simultaneous detection of calpain and Ca²⁺ in situ in order to resolve the mechanism for calpain activation in experimental uremia. We will focus also on the precise contribution of calpain and other cysteine proteases prior to and during the development of uremia, and the use of multiple cysteine protease inhibitors both in isolation and combination to prevent uremia-induced myocardial injury. A limitation of this cell culture study is that it has employed cultured myoblasts rather than primary cells because of the obvious advantages in working with immortalized cells. However, from the findings of this study, which show that calpain rises rapidly when treated with USCM, it would appear practical in the future to examine the effect of USCM treatment further by using freshly isolated rat myocytes.

To summarize, we have demonstrated that calpain activity is elevated in both in vivo and in vitro models of experimental uremia. Uremic rats had a 3.4-fold increase in calpain activity compared to control animals while lysates derived from cells incubated in uremic serum conditioned media had a fourfold higher increase in DNA fragmentation than those incubated with healthy human serum. Uremia-induced DNA fragmentation in vitro in cell lysates could be significantly reduced by treatment with two chemically distinct specific calpain inhibitors and a broad cysteine protease inhibitor, but was unaffected by caspase-3 inhibition. An in situ calpain assay revealed that significant increases in calpain activity occur within 3.5 hours in living heart derived myoblasts incubated in media enriched with uremic serum compared to those incubated with serum from healthy human volunteers. We conclude that calpain is probably a key mediator in the development uremic cardiomyopathy, but that other cysteine proteases are likely to play an important role.

References

References

1.	Raine AE, Margreiter R & Brunner FP et al. Report on management of renal failure in Europe, XXII, 1991. Nephrol Dial Transplant 1992; 7: 7–35. | PubMed | ISI |
2.	Wang KKW. Calpain and caspases: Can you tell the difference? Trends Neurosci 2000; 23: 20–26. | Article | PubMed | ISI | ChemPort |
3.	Huang Y & Wang KKW. The calpain family and human disease. Trends Mol Med 2001; 7: 355–362. | Article | PubMed | ISI | ChemPort |
4.	Thompson VF & Goll DE. Purification of -calpain, m-calpain, and calpastatin from animal tissues. Methods Mol Biol 2000; 144: 3–16. | PubMed | ChemPort |
5.	Iwamoto H, Miura T & Okamura T et al. Calpain inhibitor-1 reduces infarct size and DNA fragmentation of myocardium in ischemic/reperfused rat heart. J Cardiovasc Pharmacol 1999; 33: 580–586. | Article | PubMed | ISI | ChemPort |
6.	Tolnai S & Korecky B. Calcium-dependent proteolysis and its inhibition in the ischemic rat myocardium. Can J Cardiol 1986; 2: 42–47. | PubMed | ChemPort |
7.	McDonald MC, Mota-Filipe H & Paul A et al. Calpain inhibitor I reduces the activation of nuclear factor-B and organ injury/dysfunction in hemorrhagic shock. FASEB J 2001; 15: 171–186. | Article | PubMed | ISI | ChemPort |
8.	Liu X, Rainey JJ, Harriman JF & Schnellmann RG. Calpains mediate acute renal cell death: Role of autolysis and translocation. Am J Physiol Renal Physiol 2001; 281: F728–F738. | PubMed | ISI | ChemPort |
9.	Stern MD & Silverman HS. Ionic basis of ischaemia cardiac injury: Insights from cellular studies. Cardiovasc Res 1994; 28: 581–597. | PubMed | ISI | ChemPort |
10.	Ylitalo KV, Ala-Rämi A & Liimatta EV et al. Intracellular free calcium and mitochondrial membrane potential in ischemia/reperfusion and preconditioning. J Mol Cardiol 2000; 32: 1223–1238. | ISI | ChemPort |
11.	Borutaite V, Budriunaite A, Morkuniene R & Brown GC. Release of mitochondrial cytochrome c and activation of cytosolic caspases by myocardial ischaemia. Biochim Biophys Acta 2001; 1537: 101–109. | PubMed | ISI | ChemPort |
12.	Huang J-Q, Radinovic S, Rezaiefar P & Black SC. In vivo myocardial infarct size reduction by a caspase inhibitor administered after the onset of ischemia. Eur J Pharm 2000; 402: 139–142. | ISI | ChemPort |
13.	Amann K & Ritz E. Cardiac disease in chronic uremia: Pathophysiology. Adv Ren Replace Ther 1997; 4: 212–224. | PubMed | ChemPort |
14.	McMahon AC, Vescovo G & Dalla Libera L et al. Contractile dysfunction of isolated ventricular myocytes in experimental uraemia. Exp Nephrol 1996; 4: 144–150. | PubMed | ISI | ChemPort |
15.	Edelstein CL. Calpain activity in rat renal proximal tubules. An in vitro assay. Methods Mol Biol 2000; 144: 233–238. | PubMed | ChemPort |
16.	Sasaki T, Kikuchi T & Yumoto N et al. Comparative specificity and kinetic studies on porcine calpain I and calpain II with naturally occurring peptides and synthetic fluorogenic substrates. J Biol Chem 1984; 259: 12489–12494. | PubMed | ChemPort |
17.	Edelstein CL, Wieder ED & Yaqoob MM et al. The role of cysteine proteases in hypoxia-induced rat renal proximal tubular injury. Proc Natl Acad Sci USA 1995; 92: 7662–7666. | PubMed | ChemPort |
18.	Suzuki K, Kostin S & Person V et al. Time course of the apoptotic cascade and effects of caspase inhibitors in adult rat ventricular cardiomyocytes. J Mol Cell Cardiol 2001; 33: 983–994. | Article | PubMed | ISI | ChemPort |
19.	Yamakawa H, Banno Y & Nakashima S et al. Crucial role of calpain in hypoxic PC12 cell death: Calpain, but not caspases, mediates degradation of cytoskeletal proteins and protein kinase C- and . Neurol Res 2001; 23: 522–530. | Article | PubMed | ISI | ChemPort |
20.	Ray SK, Fidan M & Nowak MW et al. Oxidative stress and Ca2+ influx upregulate calpain and induce apoptosis in PC12 cells. Brain Res 2000; 852: 326–334. | Article | PubMed | ISI | ChemPort |
21.	Chen M, He H & Zhan S et al. Bid is cleaved by calpain to an active fragment in vitro and during myocardial ischemia/reperfusion. J Biol Chem 2001; 276: 30724–30728. | Article | PubMed | ISI | ChemPort |
22.	DeBiasi RL, Edelstein CL, Sherry B & Tyler KL. Calpain inhibition protects against virus-induced apoptotic myocardial injury. J Virol 2001; 75: 351–361. | Article | PubMed | ISI | ChemPort |
23.	Ueda N, Walker PD, Hsu S & Shah SV. Activation of a 15-kDa endonuclease in hypoxia/reoxygenation injury without morphologic features of apoptosis. Proc Natl Acad Sci USA 1995; 92: 7202–7206. | PubMed | ChemPort |
24.	Ueda N & Shah SV. Tubular cell damage in acute renal failure-apoptosis, necrosis, or both. Nephrol Dial Transplant 2000; 15: 318–323. | PubMed | ISI | ChemPort |
25.	Fujimaki S, Kubohara Y, Kobayashi I & Kojima I. Caspase-independent apoptosis induced by differentiation-inducing factor of Dictostelium discoideum in INS-1 cells. Eur J Pharm 2001; 421: 93–100. | ISI | ChemPort |
26.	Li M, Kondo T & Zhao Q-L et al. Apoptosis induced by cadmium in human lymphoma U937 cells through Ca2+-calpain and caspase-mitochondria-dependent pathways. J Biol Chem 2000; 275: 39702–39709. | Article | PubMed | ISI | ChemPort |
27.	Liang Y, Yan C & Schor NF. Apoptosis in the absence of caspase 3. Oncogene 2001; 20: 6570–6578. | Article | PubMed | ISI | ChemPort |
28.	Miyazaki K, Yoshida H & Sasaki M et al. Caspase-independent cell death and mitochondrial disruptions observed in the Apaf1-deficient cells. J Biochem 2001; 129: 963–969. | PubMed | ISI | ChemPort |
29.	Ren JG, Xia HL, Just T & Dai YR. Hydroxyl radical-induced apoptosis in human tumor cells is associated with telomere shortening but not telomerase inhibition and caspase activation. FEBS Lett 2001; 488: 123–132. | Article | PubMed | ISI | ChemPort |
30.	Wu J, He J & Sooudi S et al. Pretreatment of HL60 cells with deoxyspergualin enhances Trail-induced apoptosis independently of caspase 3 activation. Transplant Proc 2001; 33: 278. | Article | PubMed | ISI | ChemPort |
31.	Knaapen MMW, Davies MJ & De Bie M et al. Apoptotic versus autophagic cell death in heart failure. Cardiovasc Res 2001; 51: 304–312. | Article | PubMed | ISI | ChemPort |
32.	Blomgren K, Zhu C & Wang X et al. Synergistic activation of caspase-3 by m-calpain after neonatal hypoxia-ischemia. J Biol Chem 2001; 276: 10191–10198. | Article | PubMed | ISI | ChemPort |
33.	Chua BT, Guo K & Li P. Direct cleavage by the calpain-activated protease calpain can lead to inactivation of caspases. J Biol Chem 2000; 275: 5131–5135. | Article | PubMed | ISI | ChemPort |
34.	Nakagawa T & Yuan J. Cross-talk between two cysteine protease families. Activation of caspase-12 by calpain in apoptosis. J Cell Biol 2000; 150: 887–894. | Article | PubMed | ISI | ChemPort |
35.	McGinnis KM, Gnegy ME & Park YH et al. Procaspase-3 and poly(ADP)ribose polymerase (PARP) are calpain substrates. Biochem Biophys Res Commun 1999; 263: 94–99. | Article | PubMed | ISI | ChemPort |