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Keywords:

  • axonal degeneration;
  • calpain;
  • inflammation;
  • neurodegeneration

Abstract

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. Funding
  8. References
  9. Supporting Information
Thumbnail image of graphical abstract

While multiple molecular mechanisms contribute to midbrain nigrostriatal dopaminergic degeneration in Parkinson's disease (PD), the mechanism of damage in non-dopaminergic sites within the central nervous system, including the spinal cord, is not well-understood. Thus, to understand the comprehensive pathophysiology underlying this devastating disease, postmortem spinal cord tissue samples (cervical, thoracic, and lumbar segments) from patients with PD were analyzed compared to age-matched normal subjects or Alzheimer's disease for selective molecular markers of neurodegeneration and inflammation. Distal axonal degeneration, relative abundance of both sensory and motor neuron death, selective loss of ChAT+ motoneurons, reactive astrogliosis, microgliosis, increased cycloxygenase-2 (Cox-2) expression, and infiltration of T cells were observed in spinal cord of PD patients compared to normal subjects. Biochemical analyses of spinal cord tissues revealed associated inflammatory and proteolytic events (elevated levels of Cox-2, expression and activity of μ- and m-calpain, degradation of axonal neurofilament protein, and concomitantly low levels of endogenous inhibitor – calpastatin) in spinal cord of PD patients. Thus, pathologically upregulated calpain activity in spinal cords of patients with PD may contribute to inflammatory response-mediated neuronal death, leading to motor dysfunction.

We proposed calpain over-activation and calpain-calpastatin dysregulation driving in a cascade of inflammatory responses (microglial activation and T cell infiltration) and degenerative pathways culminating in axonal degeneration and neuronal death in spinal cord of Parkinson's disease patients. This may be one of the crucial mechanisms in the degenerative process.

Abbreviations used
AD

Alzheimer‘s disease

ChAT

choline acetyltransferase

deNFP

dephosphorylated neurofilament protein

GFAP

glial fibrillary acidic protein

HD

Huntington's disease

IR

immunoreactivity

LB

Lewy bodies

MND

motor neuron diseases

MS

multiple sclerosis

NeuN

neuronal specific nuclear protein

NF-L

neurofilament light

PD

Parkinson's disease

SBDP

spectrin breakdown products

SN

substantia nigra

TUNEL

terminal deoxynucleotidyl transferase recombinant–mediated dUTP nick-end labeling

Parkinson's disease (PD) is the second most common neurodegenerative disorder and is predominantly associated with the death of midbrain nigrostriatal dopaminergic neurons. The complex pathophysiology of PD also involves non-motor and non-dopaminergic symptoms that profoundly undermine the quality of life of these patients (Olanow et al. 2011). Multiple anatomically interconnected and neurotransmitter-independent regions are vulnerable in PD, and several extranigral regions have been implicated in the multistage symptomatic progression (Braak and Del Tredici 2009). The spinal cord is one such site. Conjoint lines of evidence from clinical reports and experimental studies have suggested degeneration of spinal cord in PD as outlined in recent reviews (Knaryan et al. 2011; Vivacqua et al. 2011). In addition to substantia nigra (SN) of PD brain, analysis of postmortem familial PD revealed distribution of neurofibrillary tangles and senile plaques in the cortex and enlarged axonal spheroids in the anterior horn of lumbar spinal cord (Denson and Wszolek 1995). The presence of Lewy bodies (LB) has also been noted in spinal cord degeneration in PD and other motor neuron diseases (MND) (Vivacqua et al. 2012); however, clinical overlap of PD with MND is rare (Najim al-Din et al. 1994; Trojanowski et al. 2002). Unlike MND, progression in PD is slow and insidious, non-fatal, and includes a relatively long prodromal phase prior to the appearance of motor symptoms (Hawkes et al. 2010; Siderowf and Lang 2012). Thus, there is little evidence of drastic loss of body weight or muscle wasting in PD. The role of spinal cord degeneration in PD progression is not well-understood, nor have the inflammatory response, immunohistochemical, biochemical, and neuropathological changes in postmortem PD spinal cord been studied in detail.

A common factor in many neurodegenerative diseases, including PD, is the involvement of inflammatory processes. To this end, activated glial cells, in particular activated microglia in close proximity to neurons, have been suggested to release detrimental factors that damage or kill cells (McGeer et al. 1988; Yasuda et al. 2007; Smith et al. 2012). The various factors that promote neurodegeneration are released from activated microglia, including proteases, calpain, cytokines, reactive oxygen species, and others (Smith et al. 2012). Of note, microglia activation has been demonstrated in experimental parkinsonism induced by rotenone and MPTP/MPP+ (Czlonkowska et al. 1996; Wang et al. 2006; Samantaray et al. 2007). Furthermore, acute inflammation in chronic neurodegenerative diseases like PD and others (multiple sclerosis, ischemia, stroke) is driven by matrix metalloproteases (MMPs) as common regulators (Lo et al. 2002; Lorenzl et al. 2002; Rosell and Lo 2008; Rosenberg 2009).

PD is critically associated with mild but sustained systemic mitochondrial dysfunction and aberrant intracellular Ca2+ homeostasis (Imai and Lu 2011; Lezi and Swerdlow 2012). These will likely result in dysregulation of the Ca2+-activated neutral protease, calpain, and its sole endogenous regulator, calpastatin, in PD spinal cord. This has been observed in postmortem PD brain (Mouatt-Prigent et al. 2000; Crocker et al. 2003; Samantaray et al. 2008b). Earlier findings from our laboratory confirmed calpain-mediated degeneration of spinal cord neurons in two distinct animal models of experimental PD induced by MPTP (Chera et al. 2004; Samantaray et al. 2008b) and rotenone (Samantaray et al. 2007).

The aim of this study was to confirm whether such cellular degeneration occurs in spinal cord of patients with PD. Thus, we examined selective molecular markers of degeneration in human postmortem PD spinal cord tissue in comparison to those from age-matched normal subjects and other neurological disorders without motor deficits [Alzheimer's disease (AD)] or with motor deficits [multiple sclerosis (MS), Huntington's disease (HD)]. Our results indicated that indeed, neurons, axons, and myelin are affected in the spinal cord of PD patients, and calpain was found to be a key participant in neuronal degeneration.

Materials and methods

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. Funding
  8. References
  9. Supporting Information

Postmortem spinal cord tissue

Postmortem human spinal cord tissue specimens from PD patients were obtained from National Neurological Research Specimen Bank at the West Los Angeles Veteran's Administration Medical Center; neuropathological information is summarized in the Table 1. Age-matched tissue specimens from neurologically normal subjects were obtained from Department of Pathology at Medical University of South Carolina, Charleston, SC. A part of each tissue specimen was immersed and frozen in tissue embedding media (Histo Prep; Fisher Scientific, Fair Lawn, NJ, USA); the remainder was saved and frozen at −80°C for all other assays. ARRIVE experimental guidelines were followed along with institutional approval during the course of this study.

Table 1. Human spinal cord samples used in the study
HSB No.NeuropathologyAge (years)PMI (hours)
Microscopic descriptionFinal diagnosis
  1. HSB No., Accession No. for Human Brain and Spinal Fluid Resource Center, Los Angeles, CA; AD, Alzheimer's disease; F, female; HP, hippocampus; LB, Lewy bodies; LC, locus ceruleus; M, male; PD, Parkinson's disease; PMI, post-mortem interval; SN, substantia nigra.

  2. Spinal cord tissue specimens from age-matched normal subject (n = 4) were obtained from Department of Pathology, Medical University of SC, Charleston, SC, USA.

2907Marked neuronal loss, depigmentation in SN and LC with frequent LB in the remaining neurons of LC. Rare neocortical LB in cyngulate gyrus.PDM, 8214
3605Significantly decreased neuronal cellularity, prominent associated gliosis, extracellular pigment, and a few scattered intracytoplasmic LB.PDM, 7318.5
2977Marked neuronal loss, depigmentation, and frequent LB in SN and LC.PDM, 7015.5
2898Marked neuronal loss, depigmentation, moderate reactive gliosis, and frequent LB in LC and SN. Multifocal hemorrhagic necrosis suggestive of agonal sepsis or disseminated intravascular coagulopathy.

PD

Multifocal hemorrhagic necrosis

M, 8416
3742Neuronal loss, gliosis, extracellular pigment, perivascular hemosiderin, macrophage activity, and scattered LB in SN.PDM, 7423
3220Decreased number of pigmented neurons, mild extracellular pigment, and a few scattered LB in SN and LC.PDF, 73 9.5
2286Neuronal loss resulting in a few remaining intact neuromelanin neurons in SN. No intraneuronal but extraneuronal LB in SN. Similar inclusions are found in the supraoptic nucleus neurons. Insular and temporal cortex show scattered eosinophillic cytoplasmic inclusions.

PD

Probable cortical LB dementia

No evidence of AD

F, 8115
2163Neuronal loss in SN with some pigmentary incontinence and gliosis. Neuronal loss and gliosis in HP. Probable LB in cortex.

Probable idiopathic

PD

F, 83 9

Immunofluorescent staining

Human spinal cord samples were warmed to −20°C and 5 μm thick sections were sliced using a Leica CM1850 cryostat (Leica, Deerfield, IL, USA). Tissue sections were fixed in 95% ethanol, rinsed in phosphate-buffered saline (PBS, containing 137 mM NaCl, 2.7 mM KCl, 11.9 mM phosphates, pH 7.4) and stored in the same buffer at 4°C for further studies within a week. Sections were blocked in PBS containing 2% appropriate serum (horse, goat or sheep), and then incubated with respective pri-mary IgG antibodies. Single immunofluorescent staining was performed to detect pan-neuronal neurofilament (deNFP, de-phosphorylated neurofilament protein), glial fibrillary acidic protein (GFAP), ionized calcium-binding adapter molecule 1 (Iba-1), pan-axonal neurofilament protein (NFP, SMI312), and T cell antigen receptor associated protein complex (CD3).

Terminal deoxynucleotidyl transferase recombinant–mediated dUTP nick-end labeling (TUNEL) was performed following the manufacturer's protocol (Promega, Madison, WI, USA). Sections prefixed in 95% ethanol were immersed in 4% methanol-free formaldehyde, washed in PBS, equilibrated in buffer, and incubated with digoxigenin labeled nucleotides (Roche, Indianapolis, IN, USA) and recombinant TdT enzyme (Promega) using a humidified Hybaid OmniSlide Thermal Cycler Hybaid Ltd., Teddington, UK. TUNEL reaction was terminated with 2x NaCl/Na-citrate solution, and unincorporated nucleotides were removed by three rinses in PBS. Spinal cord sections were double stained with primary antibodies, to detect co-localization of TUNEL with neuronal nuclei marker (NeuN) or motoneuronal marker choline acetyltransferase (ChAT) overnight at 4°C, and images were captured as described earlier (Samantaray et al. 2007).

Western blotting

Human spinal cord tissues (cervical and thoracic) were homogenized in an ice-cold homogenizing buffer (50 mM Tris-HCl, pH 7.4; 5 mM EGTA) with phenylmethylsulfonyl fluoride (1 mM); protein concentration was estimated with Coomassie Plus™ Protein Assay Reagent (Pierce, Rockford, IL, USA) at 595 nm. Samples were equilibrated (1 : 1 v/v) in a sample buffer (62.5 mM Tris-HCl, pH 6.8, 2% sodium dodecyl sulfate, 5 mM β-mercaptoethanol, 10% glycerol), boiled, and briefly spun before sodium dodecyl sulfate-soluble supernatant was collected. Samples were diluted to a final protein concentration of 1.5 mg/mL with mix (1 : 1 v/v) of homogenizing and sample buffers containing bromophenol blue dye (0.01%). Protein was resolved in a 4–20% pre-cast gradient gel (Bio-Rad Laboratories, Hercules, CA, USA) at 100 V for 60 min, and transferred to Immobilon™-P polyvinylidene fluoride microporous membranes (Millipore, Bedford, MA, USA). Western blotting was carried out according to Samantaray et al. (2007). Spectrin breakdown products (SBDP), neurofilament light protein (NF-L), 80 kDa μ- and m-calpain, calpastatin, and Cox-2, were detected (antibody specifications in Table S1). Calpain was detected with rabbit polyclonal antibody [1 : 500 dilution; (Banik et al. 1983; Samantaray et al. 2007)]. Blots, except those for spectrin, were re-probed for β-actin, as a protein loading control.

Semi-quantitative reverse transcription-PCR

Transcriptional expression of μ-calpain (CAPN1; 239 bp), m-calpain (CAPN2, 131 bp), and calpastatin (CAST; 188 bp), β-actin (ACTB; 198 bp), human 18S ribosomal RNA (245 bp) were analyzed following the previously described method of reverse transcription (RT)-PCR (Shields and Banik 1998; Shields et al. 1999) with modification. Total RNA was extracted from normal (n = 4) and PD (n = 8) spinal cord tissues by using TRI Reagent solution (Ambion, Applied Biosystems, Foster City, CA, USA); RNA yield was determined by measuring its absorbance at 260 nm; RNA purity was estimated by A260/A280 ratio. Generation of cDNA was performed through RT reaction in Mastercycler gradient (Eppendorf, Hamburg, Germany). The reaction mixture contained 2 μg of RNA sample and High Capacity cDNA Reverse Transcriptase Kit (Applied Biosystems), and cycles were run at 25°C (10 min), 37°C (120 min), and 85°C (5 min). Aliquots of cDNA were incubated for PCR amplification with respective primers (see Table S2). Human 18S ribosomal RNA and β-actin were amplified as internal controls. PCR amplification was performed at following thermal cycling parameters: 1 cycle at 95°C (10 min); 40 cycles at 95°C (30 s), 60–65°C (1 min), and 72°C (1 min); 1 cycle at 72°C (10 min), and held at 4°C. RT-PCR products were analyzed by resolving on Mini-PROTEAN 10% TBE precast gels (Bio-Rad) at 100 V for 2 h. Gels were incubated with SYBR Safe DNA gel stain (Molecular probes; Life Technologies, Grand Island, NY, USA) dissolved in Tris-EDTA buffer pH 8.0 for 5–10 min. PCR products were visualized in Alpha-Innotech FluorChem FC2 Imaging System (Cell Biosciences, San Jose, CA, USA) using UV transilluminator and green filter. Photomicrographs were saved and analyzed.

Statistical analysis

Photomicrographs of immunofluorescent staining were quantified for number of pixels using the NIH Image 1.63 software (NIH, Bethesda, MD, USA). Optical density values of PCR products and immunoreactivity (IR) of protein bands from western blotting were analyzed with NIH ImageJ 1.45 software. Data were analyzed by Student's t-test and/or anova and expressed as mean ± SEM. Alterations in normal and PD or control represented as percent (%) of changes (increase/decrease) were considered significant at * 0.05 compared with normal.

Results

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. Funding
  8. References
  9. Supporting Information

Neuronal death in PD spinal cord

Spinal cord tissue samples (cervical, thoracic, and lumbar) of eight PD patients and four age matched neurologically normal subjects were investigated to evaluate the extent of neuronal damage with TUNEL assays. Spinal cord sections (5 μm) were stained for TUNEL combined with NeuN or ChAT; captured images were appropriately co-localized and compared. Microscopic observation of immunofluorescent images revealed some traces of TUNEL positive IR in normal spinal cord sections whereas, significant co-localization of TUNEL and NeuN positive IR was detected in PD spinal cord (intense yellow when merged), both in dorsal and ventral regions, indicating significant neuronal degeneration in cervical and thoracic PD spinal cord (Fig. 1a). Furthermore, merged images revealed marked co-localization of TUNEL and ChAT (yellow) in ventral horns of cervical and thoracic PD spinal cord compared to normal spinal cord tissue (Fig. 1b). In contrast, neuronal death in the same areas of spinal cord of patients with AD was negligible, including ChAT+ neurons in the ventral regions compared with PD (Fig. 1). Thus, pronounced neuronal degeneration and selective motoneuronal degeneration was found in cervical and thoracic PD spinal cord compared with normal subjects or AD. Similar findings were seen in lumbar spinal cord of PD patients (data not shown).

image

Figure 1. Terminal deoxynucleotidyl transferase recombinant–mediated dUTP nick-end labeling (TUNEL) with neuronal specific nuclear protein (NeuN) and ChAT suggested neuronal degeneration is involved in motoneuronal demise in PD spinal cord. (a) Merged images of TUNEL (red) and NeuN (green) in cervical and thoracic spinal cord sections (5 μm) of neurologically normal subjects (n = 4) and PD patients (n = 12). Co-localization (yellow) of TUNEL with NeuN IR shown by arrows signify neuronal death in dorsal and ventral horns of PD spinal cord; this is absent in normal subjects. (b) Merged images of TUNEL (red) and ChAT (green) in cervical and thoracic spinal cord of normal subjects (n = 4) and PD patients (n = 12) demonstrate damaged ventral motoneurons (yellow) in PD spinal cord, which was absent in normal subjects; magnification 200x.

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Degradation of cytoskeletal protein and axonal degeneration in PD spinal cord

In PD brain, striatal dopaminergic terminals are significantly denervated. To determine whether distal axonal degeneration is present in PD spinal cord, postmortem cervical and thoracic tissues were examined for levels of 68 kDa NF-L protein. Western blotting was performed using a monoclonal antibody that specifically recognizes 68 kDa NF-L, but does not react with the other two of the triplet NF proteins (heavy or medium). Significantly reduced NF-L levels were found in cervical PD spinal cord (29%; * 0.05) compared with normal cases, while changes in thoracic tissue (by 11%) were not found to be significant (Fig. 2a and b). Along with the main 68 kDa NF-L, three distinct immunoreactive bands of lower molecular weight (presumably truncated NF-L fragments) were seen in each of cervical and thoracic samples of normal and PD specimens. Western blot analysis showed differences in the patterns of these NF-L fragments in PD versus normal tissues. A reduction in NF protein was earlier reported in PD brain (Hill et al. 1993), possibly related to accumulation of NF proteins within LB. Axonal degeneration was further confirmed by demonstrating increased levels of deNFP (de-phosphoneurofilament protein) in spinal cord sections. Extensive labeling for deNFP was detected in PD spinal cord compared with normal cases (Fig. 2c and d); values of deNFP IR (pixels) were 53% and 55% higher in cervical and thoracic sections, respectively, than in normal spinal cord; * 0.05. Taken together these findings support distal axonal degeneration in PD spinal cord.

image

Figure 2. Reduced axonal protein NF-L with elevated deNFP profiles in postmortem PD spinal cord. (a–b) Representative immunoblots and bar graphs corresponding to the 68 kDa band showed % change of NF-L IR in cervical (n = 6) and thoracic (n = 8) sections of PD spinal cord compared to neurologically normal subjects (n = 4). Reduced NF-L IR (68 kDa) was found in cervical spinal cord, whereas changes in thoracic were not significant. Of note, the monoclonal antibody against NF-L distinctly recognized three more bands in all normal samples analyzed, which were markedly reduced in both cervical and thoracic PD spinal cord tissues. As positive controls, significant loss of NF-L was seen in MS (n = 2) and HD (n = 2). (c–d) Representative photomicrographs of deNFP immunofluorescence staining and corresponding quantification of pixels indicate prominent increase of deNFP in cervical and thoracic spinal cord of PD patients (n = 12), compared with normal subjects (n = 4). Data are expressed as mean ± SEM; * 0.05 denotes statistically significant difference between spinal cord of PD patients and normal subjects; magnification 200x.

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Changes in inflammatory responses in PD spinal cord

Studies were performed to assess the inflammatory events including reactive gliosis (astrocytes and microglia) and participation of T cells in PD spinal cord compared to other neurological diseases (AD, MS, HD). Immunofluorescent staining in cervical, thoracic, and lumbar spinal cord for astroglial cell marker GFAP and microglial cell marker Iba-1 depicted presence of reactive gliosis in PD spinal cord compared with AD (data represented in lumbar spinal cord in Fig. 3a, upper and lower panels, respectively). We further spanned the Iba-1 staining in multiple PD patient spinal cord along with the pan-axonal neurofilament marker (SMI312); Iba-1 staining was found pervading in the ventral horn as well as in the white matter (Fig. 3b). Further activation of inflammatory mediator Cox-2 was detected by immunoblotting in cervical and thoracic spinal cord (Fig. 3c); Cox-2 levels were significantly increased in cervical (27%) and thoracic (30%) PD spinal cord tissue compared with normal (Fig. 3d). Immunofluorescent staining for T cell marker CD3 was also significantly greater in PD compared with normal suggesting T cell infiltration in PD spinal cord (Fig. 3e, data represented in cervical).

image

Figure 3. Associated inflammatory cascades in spinal cord degeneration in PD. (a) Marked activation of astroglial cells (GFAP) and microglial cells (Iba-1) in PD spinal cord were found via immunofluorescent staining. Images show increased GFAP IR (upper panel) and Iba-1 IR (lower panel) in lumbar PD spinal cord (n = 12) compared to AD (n = 3); profound reactive gliosis was seen in MS and HD spinal cord, represented as positive reference. (b) Double immunostaining of Iba-1 with pan-axonal NFP (SMI312) shown in ventral horn (upper panel) and white matter (lower panel) of lumbar PD spinal cord indicating presence of activated microglia in both white and gray matter. (c–d) Pro-inflammatory Cox-2 protein levels were more expressed in PD than in normal spinal cord. Immunoblots and corresponding quantification of the 72–70 kDa band showing % change of Cox-2 IR in cervical (n = 6) and thoracic (n = 8) spinal cord sections of PD patients compared to those of normal subjects (n = 4); * 0.05. (e) CD3 immunostaining showed T cell infiltration in cervical PD spinal cord (n = 8) compared to normal (n = 4).

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Dysregulation of calpain-calpastatin in PD spinal cord

Up-regulation of calpain expression/activation and dysregulation of calpain-calpastatin balance were studied to assess the underlying mechanisms of neuronal death, axonal degeneration, and inflammatory cascades in PD spinal cord. The possibility of calpain-calpastatin gene alteration was tested with semi-quantitative PCR, and calpain expression and activity were evaluated by western blotting in PD spinal cord and normal controls (Fig. 4). Profiles of calpain1, 2, and calpastatin mRNA were found unaltered in PD spinal cord as compared to normal controls (Fig. 4a and d). However, western blot analysis using a previously characterized polyclonal antibody against μ-calpain (calpain 1) and m-calpain (calpain 2) showed significant increases in both isoforms in PD spinal cord compared with normal controls. Quantitative analysis showed 50–70% and 55–65% increases in 76 kDa (active) calpain and the proenzyme (80 kDa), respectively, for both forms of calpain in cervical PD spinal cord (Fig. 4b and e). Furthermore, levels of calpastatin were found to be approximately 30% diminished in PD spinal cord compared to those in normal in cervical region. Repeating experiments with thoracic cord samples showed similar findings for calpain and calpastatin at both transcriptional and translational levels (data not presented).

image

Figure 4. Calpain and calpastatin profiles were altered at protein levels only. PD spinal cord (n = 6) relative to normals (n = 4) was tested with RT-PCR and western blotting. (a, b) Transcriptional expression of m-calpain (131 bp), and calpastatin (188 bp) in normal and PD spinal cord; β-actin (198 bp) and 18SrRNA (245 bp) were used as internal controls for each sample. Corresponding bar graphs indicated no significant alteration in transcriptional activation/repression of calpain or calpastatin in PD compared with normals (* 0.05). (c, d) Immunoblots demonstrated increased levels of 80 kDa m- and μ-calpain in PD spinal cord relative to normal subjects. The active forms of 76 kDa m- and μ-calpain were also detected in PD spinal cord, but significantly less in normal subjects. Levels of the 110 kDa calpastatin were markedly diminished in PD spinal cord relative to normal. Protein loading control was verified with 42 kDa β-actin. (e, f) SBDP immunoblots and quantified data show remarkably higher levels of 145 kDa calpain specific SBDP and 120 kDa caspase 3 specific SBDP in PD spinal cord compared with normals; * 0.05.

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These findings were further confirmed by detection of SBDP, showing increased levels of degradation of 270 kDa α-spectrin to the calpain-specific 145 kDa SBDP by 68% (Fig. 4c and f). Downstream caspase-3-specific 120 kDa SBDP was also elevated by 38% (Fig. 4c and f).

Discussion

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. Funding
  8. References
  9. Supporting Information

Salient findings in this study include significant neuronal death, axonal degeneration, and inflammatory events in spinal cord of PD patients, which were causally linked to calpain up-regulation and activity. Spinal cord involvement in PD patients has been associated with clinical symptoms including constipation, gait disturbances, and urinary/sexual dysfunction (Del Tredici and Braak 2012). Recent studies reported degeneration of spinal motoneurons in MPTP induced PD model (Samantaray et al. 2008a; Vivacqua et al. 2012). Moreover, neuronal death in SN and locus coeruleus of PD patients has been linked to increased levels of m-calpain (Mouatt-Prigent et al. 2000) and up-regulated calpain activity (Crocker et al. 2003). These findings, as well as the current results, suggest that increased calpain activity might be involved in PD spinal cord degeneration and PD pathogenesis.

A sustained neuroinflammatory response is the hallmark of several neurodegenerative diseases including PD, and the present investigation confirmed this inflammatory response in addition to neuronal damage and axonal degeneration in postmortem spinal cord of PD patients. A common early feature of the inflammatory response in all central nervous system diseases and injuries is activation of both astroglial and microglial cells. Thus, reactive gliosis is also found in conditions like MS, HD, spinal cord injury, and traumatic brain injury (McGeer et al. 1988; Shields et al. 1999; Ray and Banik 2003). Activated gliosis was reported in animal models of PD as well (Wilkin and Knott 1999; Samantaray et al. 2007). The findings of activated astroglia and microglial cells, and infiltration of CD3+ T cells in postmortem spinal cord tissue of PD patients, as reported in the current investigation, further support their involvement in the inflammatory process in this disease. The finding of CD3+ cells also suggest infiltration of T cell subtypes including CD4+ and CD8+ cells in the brain and spinal cord of PD patients as previously shown in an experimental model of PD (Laurie et al. 2007; Reynolds et al. 2007; Brochard et al. 2009). The activation of these cells has been suggested to trigger signaling to generate many pro-inflammatory factors, including reactive oxygen species, nitric oxide, cytokines (TNF-α, IL-6) and others that are detrimental to cell survival. Activated microglia have been shown to secrete inflammatory factors when in close proximity to naïve neurons, resulting in neuronal demise (Kim et al. 2005; Yasuda et al. 2007; Levesque et al. 2010; Roy et al. 2012; Smith et al. 2012). The finding of increased levels of expression and activity of Cox-2, an inflammatory marker, is closely associated with PD, and may be intricately involved in the degenerative mechanism in PD and other neurodegenerative diseases. In addition, Cox-2 activity has been suggested as a mediator of microglial activation leading to dopaminergic neuron death in experimental PD (Teismann et al. 2003; Vijitruth et al. 2006).

Calpain is known to be involved in activation of inflammatory cells (e.g., T cells, astrocytes, and microglia) and migration of immune cells; and calpain has an important role in the inflammatory process (Shields et al. 1999; Ray and Banik 2003; Butler et al. 2009) presumably in PD pathology as well. Overlapping specificities for calpain and caspase-3 has been reported in PD (Burguillos et al. 2011), experimental parkinsonism (Samantaray et al. 2007) and motoneuronal cells (VSC 4.1) exposed to parkinsonian toxins MPP+ and rotenone (Samantaray et al. 2011). In present investigation we detected increased levels of 120 kDa SBDP, which indicated caspase-3 activation in PD spinal cord compared with normal subjects. The roles of other proteases, for example, cathepsins or proteasome, in the progression of PD have also been previously studied. However, with the exception of MMPs, the results of these studies have not been definitive. MMPs have been found to act as common downstream regulators in neuroinflammation in strokes (Lo et al. 2002) and progressive neurodegenerative diseases such as PD (Rosenberg 2009). MMP-9 expression and activity in striatal degeneration was demonstrated, and treatment of PD animals with the MMP inhibitor RO-28-2653 provided neuroprotection (Lorenzl et al. 2004). Recently, an in vitro study showed the release of MMP-3 from apoptotic dopaminergic neurons, which activated microglia and acted as a mediator of interaction between apoptotic neurons and microglia (Kim et al. 2005; Choi et al. 2008). The participation of MMP-3 in microglial activation and cytokine production was also confirmed. Thus, the inflammatory process and pro-inflammatory components, for example, gliosis, elevated Cox-2, cytokines, have become targets for therapy in many neurodegenerative disorders, including PD.

Human histological studies have demonstrated neuronal loss in the intermediolateral nucleus of PD spinal cord (Wakabayashi and Takahashi 1997). Earlier findings in MPTP-induced PD mice (Chera et al. 2004; Samantaray et al. 2008a) were confirmed with demonstration of neuronal death in ventral and dorsal spinal cord of PD patients. Spinal neuronal degeneration may be involved in the abatement of norepinephrine (NE) and serotonin (5-HT) in PD. A substantial decrease in the levels of NE and 5-HT has been shown in postmortem PD tissue. Almost 40% of PD patients suffer from sensory symptoms, which may be because of loss of 5-HT input from the rubrospinal tract (Sandyk and Lacono 1987; Sage et al. 1990). The loss of NE, which modulates α–motoneurons, could be responsible for motor dysfunction in PD. Such degeneration of neurons in both ventral and dorsal spinal cord areas, as reported in the current manuscript, may play a critical role in PD spinal dysfunction. While a relatively small number of axons from the locus coeruleus that synapse in the spinal cord, modulate motor function, the loss of these neurons may ultimately affect movement in PD (Sandyk and Lacono 1987; Weil-Fugazza and Godefroy 1993; Damier et al. 1999). White matter changes, in particular degeneration of axons and myelin needed for conduction of impulses, may also be involved in dysfunction. Structural changes in axons and myelin have been shown in PD patient white matter (Yoshikawa et al. 2004; Filley 2005; Sanfilipo et al. 2006).

Axonal degeneration is evident with reduced NF-L in human tissue and increased deNFP in both human and mouse spinal cords. NF-L is one of the triplet intermediary filament proteins (heavy, medium, and light), which are the major assembly components of myelinated axons. These proteins play an essential role in structural assembly by maintaining axonal integrity, functional conductivity and axonal transport (Perrot and Eyer 2009; Shea et al. 2009). Thus, pathological alterations in axons and disorganization of axonal elements can play a role in CNS neurodegenerative disorders (Gotow 2000; Shea et al. 2009; Szaro and Strong 2010), including PD (Galvin et al. 1999). For example, NF-L deficient mice lose 20% of motor axons, and demonstrate a reduction of axonal caliber in those remaining (Zhu et al. 1997). Depletion of the NF-L protein causes mild sensorimotor dysfunctions and spatial deficits in NF-L −/− mice, without overt signs of paresis (Dubois et al. 2005). NF-L 68 kDa protein is a calpain substrate (Pollanen et al. 1992); it was detected in cortical LBs, a morphological hallmark of PD. In contrast to normal NF phosphorylation patterns (Sternberger and Sternberger 1983), NF proteins in LBs are present in abnormally phosphorylated forms (Forno et al. 1986), where they undergo degradation through proteolysis inside ubiquitinated LBs (Galloway et al. 1988). This study showed an extensive deNFP IR detected in cervical and thoracic PD spinal cord compared to normal, indicating distal axonal degeneration in PD. Therefore, axonal degeneration (with degradation of filament proteins) by increased calpain activity in PD spinal cord may have a profound effect on motor function. Moreover, calpain inhibition has been found to improve behavioral function in MPTP-induced PD animals (Crocker et al. 2003).

Sporadic PD does not originate in the spinal cord, but spinal cord lesions are seen in all PD cases between neuropathological stages 2–6 for PD-related synucleinopathy (Del Tredici and Braak 2012). A direct correlation of spinal cord degeneration with the disease severity may not be deduced from this study. Nonetheless, spinal cord pathology in PD is more than a mere secondary phenomena; PD pathology is complex and existing dopamine replenishment therapy does not prevent the disease progression. Overall, this study provides direct evidence of neuronal degeneration, distal axonal damage, inflammation, and calpain-calpastatin dysregulation in PD spinal cord, which in correlation with earlier reports on calpain-mediated midbrain neuronal degeneration in PD, strongly corroborate a crucial role of calpain in the complex pathogenesis of PD.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. Funding
  8. References
  9. Supporting Information

The authors thank Dr. Wallace W. Tourtellotte in National Neurological Research Specimen Bank at the West Los Angeles Veteran's Administration Medical Center and Dr. Cynthia Wahl in Department of Pathology, Medical University of South Carolina for their valuable support in providing us with post-mortem spinal cord tissue for this study. Authors have no conflict of interest to declare.

Funding

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. Funding
  8. References
  9. Supporting Information

This study was funded in part by the R01 grants from National Institute of Neurological Disorders and Stroke of the National Institutes of Health (NINDS-NIH; NS-62327-01A2; NS-56176 and NS-65456) and the Veterans Administration (I01 BX001262).

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  6. Acknowledgements
  7. Funding
  8. References
  9. Supporting Information
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Supporting Information

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. Funding
  8. References
  9. Supporting Information
FilenameFormatSizeDescription
jnc12374-sup-0001-TableS1.docWord document134KTable S1. Primary antibodies used in the study.
jnc12374-sup-0002-TableS2.docWord document99KTable S2. Human oligonucleotide primers used in the study.

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