The role of aberrant mitochondrial bioenergetics in diabetic neuropathy

Neurobiology of Disease 51 (2013) 56–65

Contents lists available at SciVerse ScienceDirect

Neurobiology of Disease journal homepage: www.elsevier.com/locate/ynbdi

Review

The role of aberrant mitochondrial bioenergetics in diabetic neuropathy Subir K. Roy Chowdhury a, Darrell R. Smith a, Paul Fernyhough a, b,⁎ a b

Division of Neurodegenerative Disorders, St Boniface Hospital Research Centre, Winnipeg, MB, Canada Department of Pharmacology & Therapeutics, University of Manitoba, Winnipeg, MB, Canada

a r t i c l e

i n f o

Article history: Received 7 November 2011 Revised 22 February 2012 Accepted 1 March 2012 Available online 9 March 2012 Keywords: AMP-activated protein kinase Axonal plasticity Axonopathy Diabetes Dorsal root ganglia Mitochondrial depolarization Nutrient excess Respiration Respiratory chain Schwann cell

a b s t r a c t Diabetic neuropathy is a neurological complication of diabetes that causes significant morbidity and, because of the obesity-driven rise in incidence of type 2 diabetes, is becoming a major international health problem. Mitochondrial phenotype is abnormal in sensory neurons in diabetes and may contribute to the etiology of diabetic neuropathy where a distal dying-back neurodegenerative process is a key component contributing to fiber loss. This review summarizes the major features of mitochondrial dysfunction in neurons and Schwann cells in human diabetic patients and in experimental animal models (primarily exhibiting type 1 diabetes). This article attempts to relate these findings to the development of critical neuropathological hallmarks of the disease. Recent work reveals that hyperglycemia in diabetes triggers nutrient excess in neurons that, in turn, mediates a phenotypic change in mitochondrial biology through alteration of the AMP-activated protein kinase (AMPK)/peroxisome proliferator-activated receptor γ coactivator-1α (PGC1α) signaling axis. This vital energy sensing metabolic pathway modulates mitochondrial function, biogenesis and regeneration. The bioenergetic phenotype of mitochondria in diabetic neurons is aberrant due to deleterious alterations in expression and activity of respiratory chain components as a direct consequence of abnormal AMPK/PGC-1α signaling. Utilization of innovative respirometry equipment to analyze mitochondrial function of cultured adult sensory neurons from diabetic rodents shows that the outcome for cellular bioenergetics is a reduced adaptability to fluctuations in ATP demand. The diabetes-induced maladaptive process is hypothesized to result in exhaustion of the ATP supply in the distal nerve compartment and induction of nerve fiber dissolution. The role of mitochondrial dysfunction in the etiology of diabetic neuropathy is compared with other types of neuropathy with a distal dying-back pathology such as Friedreich ataxia, Charcot–Marie–Tooth disease type 2 and human immunodeficiency virus-associated distal-symmetric neuropathy. © 2012 Elsevier Inc. All rights reserved.

Contents Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Clinical impact of diabetic neuropathy . . . . . . . . . . . . . . . . . . . . . . . . . . Nerve pathology in diabetic neuropathy . . . . . . . . . . . . . . . . . . . . . . . . . Mitochondrial ultrastructure in diabetic neuropathy . . . . . . . . . . . . . . . . . . . Neuronal mitochondrial inner membrane polarization status is diminished in diabetes . . . Aberrant response to mitochondrial inner membrane hyperpolarization in diabetic neurons . Mitochondrial proteome is altered in DRG from diabetic rats . . . . . . . . . . . . . . . Nutrient excess and depressed AMPK signaling . . . . . . . . . . . . . . . . . . . . . . Studying mitochondrial function in cultured neurons . . . . . . . . . . . . . . . . . . . The cellular bioenergetic profile exhibits reduced adaptability in diabetic neurons . . . . . Mitochondrial dysfunction in other neuropathic diseases . . . . . . . . . . . . . . . . . CMT2. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . HSV-DSP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Friedreich ataxia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

⁎ Corresponding author at: R4046-351 Taché Ave, St Boniface Hospital Research Centre, Winnipeg, Canada MB R2H 2AL. Fax: + 1 204 239 4092. E-mail address: [email protected] (P. Fernyhough). Available online on ScienceDirect (www.sciencedirect.com). 0969-9961/$ – see front matter © 2012 Elsevier Inc. All rights reserved. doi:10.1016/j.nbd.2012.03.016

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

57 57 57 57 58 59 59 59 59 60 60 60 61 61

S.K.R. Chowdhury et al. / Neurobiology of Disease 51 (2013) 56–65

Schwann cell mitochondria involvement in axonal degeneration in diabetes . . . . . . . Putative mechanism(s) of distal axonopathy linked to neuronal mitochondrial dysfunction Conclusion and the future . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Introduction This review will focus on aspects of mitochondrial dysfunction in neurons and Schwann cells in the setting of diabetes and diabetic neuropathy. Diabetic neuropathy, a common neurological complication of diabetes, exhibits a distal dying-back process of nerve fiber degeneration that is seen in other neuropathic diseases such as Friedreich ataxia, Charcot–Marie–Tooth disease type 2 (CMT2; an axonal form) and human immunodeficiency virus-associated distalsymmetric neuropathy (HIV-DSP). Importantly, it is now clear that all of these diseases incorporate a component involving mitochondrial dysfunction. In other neuropathies with a dying-back type axonopathy, such as leprosy and myriad chemical-induced neuropathies, the specific role of mitochondrial dysfunction in etiology is less well characterized. However, the reader is referred to work by Bennett et al. with regard to mitochondrial dysfunction in chemotherapy (taxol or oxaliplatin)-induced peripheral neuropathy (Bennett et al., 2011; Xiao et al., 2011; Zheng et al., 2011). This review will concentrate on newly characterized features of mitochondrial physiology that are abnormal in diabetic neuropathy and will take an axon-centric view but will also consider Schwann cell involvement, although data in this area are lacking. For background information the reader is directed toward recent exhaustive reviews covering broad aspects of the putative etiologies underpinning neuropathy and the diabetic complications (Calcutt et al., 2009a; Sugimoto et al., 2008; Tomlinson and Gardiner, 2008) and more specifically the role of mitochondrial dysfunction in the development of diabetic complications (Sivitz and Yorek, 2010). Dyslipidemia has also been introduced as an etiological factor contributing to diabetic neuropathy (Vincent et al., 2009), although supporting experimental data is limited and awaits verification. For more specific coverage of mitochondrial abnormalities in diabetic neuropathy, with a focus on metabolic pathway involvement and alterations in organelle biogenesis and trafficking, the reader is referred to the following articles (Chowdhury et al., 2011; Fernyhough et al., 2010).

Clinical impact of diabetic neuropathy The World Health Organization (WHO) predicts that by 2025 there will be 300 million people with diabetes. In North America 19 million people currently have diabetes with an incidence of 6% and rising. Of those, approximately 90–95% has non-insulindependent diabetes mellitus (type 2 diabetes) and 5–10% has insulin-dependent diabetes mellitus (type 1 diabetes). In the USA approximately US$25 billion per annum is spent on the treatment of diabetic complications that include retinopathy, nephropathy, heart disease and neuropathy. In 1998 approximately US$15 billion of the health service expenditure was associated with neurological complications (sensory and autonomic neuropathy and blindness). The incidence of symmetrical polyneuropathy, the most common presentation of neuropathy, in diabetic patients can be 50% or higher and leads to incapacitating pain, sensory loss, foot ulceration (up to 2 million Americans have this complaint), infection, gangrene and poor wound healing. Lower extremity amputation often follows and accounts for approximately 80,000 cases each year in the USA. There is no effective therapy for diabetic symmetrical polyneuropathy or any form of neuropathy, only palliative treatment is available at

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

57

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

61 62 63 63 63

the present time. These alarming figures are predicted to rise by approximately 5-fold over the next 10 years due to the epidemic in obesity and the associated increase in incidence and earlier time of onset of type 2 diabetes (Fernyhough et al., 2010). Nerve pathology in diabetic neuropathy Symmetrical polyneuropathy in types 1 and 2 diabetic patients is associated with a reduction in motor and sensory nerve conduction velocity and appearance of structural changes in the peripheral nerves including endoneurial microangiopathy (Malik et al., 2005), abnormal Schwann cell pathology (Kalichman et al., 1998), axonal degeneration, paranodal demyelination, and loss of myelinated and unmyelinated fibers (Said, 2007; Yagihashi, 1996); the latter due to a dying-back of distal axons that presents clinically as reduced epidermal nerve fiber density (Ebenezer et al., 2011; Kennedy et al., 1996; Quattrini et al., 2007). Neurodegeneration is most profound in the longest axons of neurons (Said, 2007), and defective axon regeneration impedes tissue re-innervation (Ebenezer et al., 2011; Polydefkis et al., 2004). These clinical endpoints as well as indications of pain are observed in animal models with diabetic neuropathy, with demyelination being the only pathological sign that is not present reproducibly (Beiswenger et al., 2008; Christianson et al., 2003, 2007; Francis et al., 2009; Jolivalt et al., 2008; Mizisin et al., 2007). While demyelination is not present in some animal models, in long term streptozotocin (STZ)-induced diabetic mice (a model of type 1 diabetes) there was evidence of myelin thinning (Kennedy and Zochodne, 2005a). Furthermore, a recent gene array study of nerve in db/db diabetic mice (a model of type 2 diabetes) revealed a range of deficits in myelin protein-associated gene expression (Pande et al., 2011). Complementary studies using co-cultures of sensory neurons and Schwann cells and the examination of STZ-diabetic mice suggest demyelination, and the attendant deficits in sensory function may be associated with hyperglycemia-induced aberrant caveolin and neuregulin/erbB2 signaling (Dobrowsky et al., 2005; McGuire et al., 2009; Yu et al., 2008). Shrinkage of perikaryal volume occurs within the dorsal root ganglia (DRG) in animal models of diabetes, however, there is no molecular signature or sign of apoptosis-dependent loss of sensory neuron perikarya in diabetic humans or animals (Burnand et al., 2004; Cheng and Zochodne, 2003; Kamiya et al., 2006; Schmidt et al., 1997; Sidenius and Jakobsen, 1980). Nevertheless, loss of small neurons, through a yet to be defined mechanism, does occur in mouse and rat models (Jiang et al., 2004; Kamiya et al., 2006; Kennedy and Zochodne, 2005a). The distal dying-back and formation of axonal dystrophy (with swellings) of axons are critical pathological features (Kennedy et al., 1996; Lauria et al., 2003; Polydefkis et al., 2004; Schmidt, 2002; Schmidt et al., 1981; Zherebitskaya et al., 2009) and mimic axonal pruning and degeneration observed in the CNS and PNS in other pathological states (Nja and Purves, 1978; Verkhratsky and Fernyhough, 2008). Mitochondrial ultrastructure in diabetic neuropathy During peripheral nerve degeneration in diabetes in both humans and animal models subtle changes in mitochondrial number and size have been described in Schwann cells of myelinated and non-

58

S.K.R. Chowdhury et al. / Neurobiology of Disease 51 (2013) 56–65

myelinated axons, and glycogen accumulation in the outer compartment of the mitochondrion was observed in axons (Kalichman et al., 1998). The Schwann cells exhibited degenerative changes including enlargement of mitochondria and effacement of cristae, however, axonal mitochondria appeared relatively normal (Kalichman et al., 1998). In the peripheral nerve in animal models a similar structural phenotype of Schwann cells and mitochondria was observed compared with humans, although in the peroneal nerve in a feline model of diabetes there was evidence of more severe degenerative axonal pathology; namely, enlarged mitochondria (Kalichman et al., 1998; Mizisin et al., 2007). In long term Bio-Breeding Wistar (BB/Wistar) diabetic rats (a model of type 1 diabetes) the ultrastructure of mitochondria in neuronal cell bodies within the DRG was reportedly unremarkable (Kamiya et al., 2006). In 24 week db/db diabetic mice the numbers of mitochondria were amplified in myelinated and unmyelinated axons within the dorsal root (Vincent et al., 2010). Mitochondria are a normal part of presynaptic boutons, likely reflecting their metabolic activity, but have been recently found in increased numbers in autophagic vacuoles admixed with synaptic vesicles in murine diabetic sympathetic ganglia (Schmidt et al., 2008). More impressive are the accumulated mitochondria, tightly aggregated without a significant amount of intervening cytoplasm, in post-synaptic dendrites in which they may produce nearly pure aggregates often composed of smaller mitochondria than those in adjacent cell bodies (Schmidt et al., 2009, 2011) and reviewed by Fernyhough et al. (2010). Recent studies of prevertebral sympathetic ganglia of nonobese diabetic (NOD) mice (a model of type 1 diabetes), STZtreated mice and STZ-treated NOD/severe combined immunodeficiency (SCID) mice (Schmidt et al., 2003) and in the spontaneously genetically diabetic Akita mouse (a model of type 1 diabetes) (Schmidt et al., 2009), have demonstrated the presence of striking abnormalities in mitochondrial ultrastructure. In particular, the accumulation of small, hyperchromatic dense mitochondria that were significantly smaller in magnitude than those in nondiabetic control mouse ganglia. Thus in sensory fibers of the dorsal root and in sympathetic ganglia (in perikarya and dendrites) of these animal models there is accumulation of small mitochondria and could be associated with diabetes-induced alterations in mitochondrial biogenesis and/ or trafficking (Fernyhough et al., 2010; Schmidt et al., 2008, 2009; Vincent et al., 2010). This is in contrast to the majority of findings in perikarya and afferent projections of sensory neurons in animals and humans with diabetes where the ultrastructure of mitochondria appears relatively conventional (Kalichman et al., 1998; Kamiya et al., 2006; Schmidt et al., 1997). Neuronal mitochondrial inner membrane polarization status is diminished in diabetes The ultrastructural evidence pointing to mitochondrial dysfunction (Fernyhough et al., 2010) was the primary indicator of a putative role in etiology of diabetic neuropathy until studies by Srinivasan et al. and our own work showed that in the perikarya of adult sensory neurons from STZ-diabetic rats the mitochondrial inner membrane potential was depolarized (Huang et al., 2003; Huang et al., 2005a; Srinivasan et al., 2000). Mitochondrial depolarization in STZdiabetes could be prevented by systemic treatment with low dose of insulin, that did not impact on hyperglycemia, or neurotrophin-3 (NT-3) (Huang et al., 2003; Huang et al., 2005a). Insulin and other neurotrophic growth factors could directly modulate mitochondrial polarization through a phosphoinositide 3-kinase dependent pathway in cultured neurons (Huang et al., 2005b). Fig. 1 shows that these abnormalities in mitochondrial inner membrane potential are also present in axons of sensory neurons derived from STZ-diabetic rats (Akude et al., 2011). This work has been described in more detail elsewhere (Akude et al., 2011). Adult sensory neurons were cultured for 1 day from age matched control and

Fig. 1. Mitochondria exhibit abnormal physiology in axons of sensory neurons derived from diabetic rats. (A) Shows images of fluorescence confocal microscopy using TMRM in live cultures of DRG neurons isolated from control adult rats showing effect of antimycin A and oligomycin. (B) Trace of TMRM fluorescence signal in the axons of cultured DRG neurons isolated from age matched controls (filled gray diamond) and STZ-diabetic rats (filled black diamond). (C) Shows the area under the TMRM fluorescence trace (AUC) for control (gray bar) and diabetic (black bar) neurons. The AUC was estimated from the baseline (at the point of injection) to a fluorescence level of 0.2 and between time points of 1.0 min and 6 min using sums-of-squares (shown by dotted line). Values are the means ± SEM, n = 65–80 axons; * p b 0.001 compared to control, t-test. The TMRM trace was characterized by non-linear regression (one phase exponential decay). The rate constant of decay (K) = 0.013± 0.0004 (control) and 0.006± 0.0001 (diabetic). Half-life of decay = 54.19 s (control) and 108.7 s (diabetic). The F (Fisher parametric)ratio = 409.5, P b 0.0001, control vs diabetic. The F-ratio compares the goodness-of-fit of the two curves. The red arrow indicates point of injection of antimycin A + oligomycin. (D) TMRM fluorescence trace of oligomycin-induced mitochondrial inner membrane hyperpolarization in the axons of control (gray diamond) and diabetic (black triangle) neurons. Values are the mean ± SEM, n = 65–85 axons. Inset shows the area under the TMRM fluorescence trace (AUC) for control (gray bar) and diabetic (black bar) neurons. The AUC was estimated from the baseline (at the point of injection, dotted line), and between time points of 1.0 min and 4 min using sums-of-squares. Values are the means± SEM, n = 65–80 axons,* P b 0.01 compared to control, t-test. The red arrow indicates point of injection of oligomycin. Copyright 2010 American Diabetes Association. From Diabetes, vol. 60, 2011; 288. Reprinted with permission from the American Diabetes Association.

22 week STZ-diabetic rats and loaded with tetramethylrhodamine methyl ester (TMRM) as previously described (Akude et al., 2011; Zherebitskaya et al., 2009). This dye was used at a sub-quench concentration where a decrease in fluorescence signal intensity indicated lowered mitochondrial inner membrane potential (Nicholls, 2006). The live neurons were exposed to a combination of antimycin A (inhibitor of complex III) and oligomycin (inhibitor of ATP synthase) and the fluorescence signal in axons detected by confocal microscopy.

S.K.R. Chowdhury et al. / Neurobiology of Disease 51 (2013) 56–65

Antimycin A blocks electron transfer leading to mitochondrial depolarization whereas oligomycin inhibits the ATPase and prevents reverse pumping of protons and associated generation of a proton gradient. Therefore, the mitochondrial membrane potential (and associated proton gradient) is rapidly and completely dissipated in the presence of both these drugs. In the presence of antimycin A + oligomycin, the rate of mitochondrial depolarization was more rapid in axons of normal neurons compared with diabetic neurons (Figs. 1A– C). This suggests that prior to addition of antimycin A + oligomycin, the axonal mitochondria were more highly polarized in the normal neurons compared with the diabetic neurons. Aberrant response to mitochondrial inner membrane hyperpolarization in diabetic neurons Mitochondrial physiology was further investigated in Fig. 1D by treating cultured neurons from control and diabetic rats with oligomycin alone (Akude et al., 2011). Blockade of the ATPase results in a build-up of the transmembrane proton gradient and induces hyperpolarization of the mitochondrial inner membrane as indicated by elevated TMRM fluorescence (Jekabsons and Nicholls, 2004). In normal neurons a transient hyperpolarization was observed followed by a recovery due to adaption of the respiratory chain. For example, under a high inner membrane electrochemical gradient respiratory chainderived superoxide radicals activate uncoupling proteins which then dissipate the proton gradient (Azzu et al., 2008; Echtay et al., 2003; Murphy et al., 2003). Diabetic neurons exhibited a significantly greater level of hyperpolarization upon oligomycin application and the adaptive response was impaired. This aberrant response in mitochondria of diabetic neurons could be the result of lower respiratory chain-dependent ROS production (Akude et al., 2011), deficient lipid peroxidation and/or sub-optimal uncoupling protein expression (Vincent et al., 2004a). Mitochondrial proteome is altered in DRG from diabetic rats Our recent published results demonstrating deficits in mitochondrial respiratory function in DRG of STZ-diabetic rats (Akude et al., 2011; Chowdhury et al., 2010) are in general agreement with findings described in skeletal muscle and cardiac tissue (reviewed by (Bugger and Abel, 2010; Chowdhury et al., 2011; Szendroedi et al., 2011)). For example, gene array studies in skeletal muscle from type 2 diabetic patients reveal diminished expression of oxidative phosphorylation genes (Patti et al., 2003). There was also a decreased expression of the transcriptional regulator, nuclear respiratory factor 1 (NRF-1) and the translational co-activator, peroxisome proliferator-activated receptor γ coactivator-1α (PGC-1α) (Mootha et al., 2003). Investigators have begun to use proteomic and gene array techniques in animal models to identify alterations in gene expression that presumably underpin such changes in mitochondrial physiology (Bugger et al., 2009; Pande et al., 2011; Zhang et al., 2010). In cardiac tissue of Akita mice TCA cycle proteins were down-regulated and corresponded with decreased transcripts for PGC-1α and genes of oxidative phosphorylation (Bugger et al., 2009). In cultured Schwann cells exposed to high [glucose] the proteins associated with mitochondrial dysfunction, tricarboxylic acid (TCA) cycle and oxidative phosphorylation were enhanced (Zhang et al., 2010). Microarray work on sciatic nerve of db/db mice revealed mRNA for TCA cycle proteins to be raised and transcripts for myelin proteins, axonal regeneration and neurotrophin signaling pathways to be depressed (Pande et al., 2011). The stable isotope labeling with amino acids in cell culture (SILAC)-based quantitative proteomics analysis of mitochondrial protein expression in DRG from 22 week STZ-diabetic rats by our group revealed that diabetes altered the levels of an array of proteins (mitochondrial complexes — Complex I–V, TCA cycle, oxidative stress) associated with mitochondrial function with down-regulation of

59

expression most common (Akude et al., 2011; Chowdhury et al., 2011) and corresponding most closely with findings in cardiac tissue (Bugger et al., 2009). The work supports the premise that reduced activity of the mitochondrial respiratory chain in DRG neurons could result from a proteome alteration leading to sub-optimal expression/ activity of a range of mitochondrial components.

Nutrient excess and depressed AMPK signaling The AMP-activated protein kinase (AMPK) and PGC-1α signaling axis senses the metabolic demands of cells and regulates mitochondrial function (Canto and Auwerx, 2009; Dominy et al., 2010; Feige and Auwerx, 2007; Hardie, 2008; Puigserver et al., 1998; Rodgers et al., 2008). Studies in skeletal muscle, liver and cardiac tissue have shown that the activity of AMPK and PGC-1α signaling axis declines under diabetic conditions (Bugger and Abel, 2010; Mootha et al., 2003; Patti et al., 2003; Richardson et al., 2005; Szendroedi et al., 2011). We have tested the hypothesis that deficits in AMPK/PGC-1α signaling in sensory neurons underlie compromised axonal plasticity, sub-optimal mitochondrial function and development of neuropathy in rodent models of type 1 diabetes. Phosphorylation and expression of AMPK/PGC-1α and mitochondrial respiratory chain complex proteins were down-regulated in DRG of STZ-diabetic rodents (Chowdhury et al., 2011). Recent work demonstrates that adenoviral-mediated manipulation of endogenous AMPK activity using mutant proteins modulated neurotrophin-directed neurite outgrowth in cultures of sensory neurons derived from adult rats. Addition of resveratrol, an activator of AMPK, to cultures of sensory neurons derived from rats after 3–5 months of STZ-induced diabetes significantly elevated AMPK levels, enhanced neurite outgrowth and normalized mitochondrial inner membrane polarization in axons. Treatment of diabetic rats with resveratrol normalized deficits in AMPK signaling and prevented sensory loss (Roy Chowdhury et al., in press). These data suggest that the development of distal axon loss in sensory neuropathy is linked to nutrient excess and mitochondrial dysfunction via defective signaling of the AMPK/PGC-1α pathway and has recently been reviewed (Chowdhury et al., 2011).

Studying mitochondrial function in cultured neurons With the introduction of the Seahorse Bioscience XF24 Analyzer it is now feasible to study neuronal bioenergetics in cultured neurons. The XF24 Analyzer creates a transient 7 μl chamber in specialized 24-well microplates that permits oxygen consumption rate (OCR) to be monitored in real time (Hill et al., 2009). In the studies described in Fig. 2 adult sensory neurons derived from age matched control or 4–5 month STZ-induced diabetic rats were cultured for 24 h as previously described (Akude et al., 2011; Zherebitskaya et al., 2009). The results presented duplicate recently published work (Roy Chowdhury et al., in press). Similar results were seen with neurons from STZ-diabetic mice (data not shown). The basal level of oxygen consumption, the amount of oxygen consumption linked to ATP production, the level of non-ATP-linked oxygen consumption (proton leak), the maximal respiration capacity and the non-mitochondrial oxygen consumption were determined as previously described (Brand and Nicholls, 2011; Hill et al., 2009). Oligomycin inhibits the ATP synthase leading to a build-up of the proton gradient that inhibits electron flux and reveals the state of coupling efficiency. Uncoupling of the respiratory chain by FCCP injection reveals the maximal capacity to reduce oxygen. Finally, rotenone + antimycin A were injected to inhibit the flux of electrons through complexes I and III, and thus no further oxygen was consumed at cytochrome c oxidase. The remaining OCR determined after this intervention is primarily nonmitochondrial in origin.

60

S.K.R. Chowdhury et al. / Neurobiology of Disease 51 (2013) 56–65

mitochondrial mass in sensory neuron perikarya or afferent fibers in diabetic animals or humans (Kalichman et al., 1998; Kamiya et al., 2006; Mizisin et al., 2007; Schmidt et al., 1997). Secondly, the yield of purified mitochondria from DRG is the same from age matched control versus diabetic (unpublished data). Thirdly, in our Seahorse XF24 studies the staining of cultures at study end with Mitotracker far red, which enters and binds to mitochondria irrespective of polarization state (and is well retained after fixation), revealed mitochondrial mass to be equivalent between age matched control and diabetic cultures (data not shown). Finally, complementary support for this contention derives from inspection of the real time imaging studies with TMRM in Fig. 1 that reveal alterations in mitochondrial physiology that are independent of mitochondrial number. Therefore, the deficits in OCR are due to intrinsic abnormalities in mitochondrial performance, e.g. suboptimal respiratory chain activity, as opposed to just loss of mitochondria on a per cell basis. At this juncture no other work has been published that provides any additional information on the impact of diabetes on mitochondrial bioenergetics of sensory neurons. Therefore, in sensory neurons maintained in vitro, we propose that the carefully titrated uncoupled rates of mitochondrial electron flux reported the maximum activity of respiratory chain function and substrate oxidation independently of effects of variation in mitochondrial mass. Measurements of OCR in the presence of uncoupler revealed that the maximal electron transport capacity was significantly depressed in sensory neurons from STZ-diabetic rats. Reduced spare respiratory capacity, especially in neurons that have variable ATP demands, limits their ability to meet energetic needs and renders the cells more susceptible to secondary stressors (Brand and Nicholls, 2011; Hill et al., 2009). Diabetes dependent blunting of the maximal OCR and lowering of spare respiratory capacity suggests that the cells were energetically stressed and that mitochondrial workload was increased. Mitochondrial dysfunction in other neuropathic diseases

Fig. 2. Measurement of mitochondrial function in cultured neurons using the XF24 analyzer. OCR was measured at basal level with the subsequent and sequential addition of oligomycin (1 μM), FCCP (0.1–0.75 μM), and rotenone (1 μM) + antimycin A (AA; 1 μM) to DRG neurons cultured from (A) age-matched control and (B) 4 month STZdiabetic rats. Levels of OCR are normalized per 1000 cells. Dotted lines, a–d in (A) have been used later for quantification of bioenergetic parameters. The OCR measurements in (A) and (B) at the 0.75 μM concentration of FCCP were plotted in (C) as % related to the basal respiration. (D) Coupling efficiency (c/a), (E) respiratory control ratio (d/b) and (F) spare respiratory capacity (d–a) were calculated after subtracting the non-mitochondrial respiration (e) as described (Brand and Nicholls, 2011). In (G) and (H) Pie charts illustrate the parameters of mitochondrial function (spare respiratory capacity (d–a), ATP-linked O2 consumption (a–b), proton leak (a–c), and non-mitochondrial O2 consumption (e)) expressed as % of maximal OCR from age-matched control and STZ-diabetic rats. Values are mean ± SEM of n = 5 replicate cultures; * p b 0.05 vs. diabetic (Student's t-test).

The cellular bioenergetic profile exhibits reduced adaptability in diabetic neurons OCR induced by the uncoupler FCCP (0.3–0.75 μM) in cultured neurons from diabetic rats (Fig. 2) as well as % related to the basal respiration was significantly decreased compared to neurons from agematched control rats indicating impairment of maximal respiratory chain activity in the diabetic state. Spare respiratory capacity was significantly depressed in cultured neurons from diabetic vs control rats but coupling efficiency was slightly higher in neurons from diabetic rats. Pie charts are presented summarizing the different processes contributing to oxygen consumption in the cultures. These results are unlikely to be the result of any significant diabetes-induced depression in mitochondrial number or mass per cell. Firstly, there is no significant ultrastructural evidence to support increased

It is informative to compare and contrast the salient features of mitochondrial dysfunction in selective examples of neurodegenerative diseases that exhibit a distal dying-back type of neuropathy. CMT2 In Charcot–Marie–Tooth disease type 2 (CMT2) a distal dying-back axonal degeneration is predominant and 19–33% of cases has been linked to mutations in the GTPase and mitochondrial fusion protein, mitofusin 2 (MFN2) (Zuchner et al., 2004). In a series of studies the groups of Baloh and Milbrandt demonstrated that mutant MFN2 overexpressed in sensory neurons resulted in a distal dying-back neuropathy in mouse models that was characterized by compromised axonal trafficking of mitochondria (Baloh et al., 2007). The bioenergetic properties of mitochondria were not significantly altered and the role of MFN2 in mediating transport was deemed to be discrete from regulation of mitochondrial fusion (Baloh et al., 2007; Misko et al., 2010). At this juncture in diabetic neuropathy there is no clear evidence of altered trafficking of mitochondria in peripheral nerve although accumulation of small mitochondria in sensory axons in dorsal root and in sympathetic ganglia, perikarya and dendrites, implies a deficit in this regard (Schmidt et al., 2008; Schmidt et al., 2009; Vincent et al., 2010). See Fernyhough et al. for a more complete discussion on this aspect (Fernyhough et al., 2010). Studies on rates of fast axonal transport of neuropeptides and enzymes reveal no deficit in animal models of diabetes (see references below). Since the early 1990s numerous papers investigating peripheral nerve function in STZ-diabetic and BB/Wistar diabetic rats have shown axonal transport defects to be due to altered levels of cargoes. The former could be caused by reduced perikaryal neurotransmitter or enzyme synthesis or reduced receptor-mediated

S.K.R. Chowdhury et al. / Neurobiology of Disease 51 (2013) 56–65

uptake of growth factors. Several papers show that impaired axonal transport of proteins is observed while the actual rates of the axonal transport process are not affected (see Abbate et al., 1991; Calcutt et al., 1990; Delcroix et al., 1998; Hellweg et al., 1994; Kilgour et al., 1990; Schmidt et al., 1987). However, all these studies have focused on measurements of dorsal/ventral root, mid-sciatic or mesenteric nerve and detailed analysis of mitochondrial trafficking and/or vesicular transport in distal axons of afferent fibers has not been attempted, as yet, in the diabetic setting. HSV-DSP Post-mortem peripheral nerve samples from patients with HSVDSP reveal increased mutation in mitochondrial DNA (mutation mtDNA 4977) that was associated with deficits in mitochondrial protein expression (Lehmann et al., 2011). These insufficiencies were more pronounced at a distal level compared with more proximal nerve segments. Simian immunodeficiency virus (SIV) infected macaques exhibited similar abnormalities and, in addition, markers of mitochondrial function were aberrant combined with an elevation in mitochondrial-dependent reactive oxygen species (ROS) production (Lehmann et al., 2011). Studies with cultured human DRG treated with supernatants from HIV-infected macrophages also reveal mitochondrial dysfunction (membrane depolarization) with signs of oxidative stress in perikarya but not in axons (Hahn et al., 2008). Where mitochondrial dysfunction impacts the etiology of the virusinduced neurodegenerative process remains unclear; for example, does oxidative stress cause the mitochondrial dysfunction or vice versa? However, the alterations in mitochondrial phenotype, e.g. inner membrane depolarization and depressed expression of proteins, mimic the defects seen in diabetic neuropathy. Friedreich ataxia This is an autosomal recessive neurodegenerative disease induced by a GAA repeat expansion in intron 1 of the frataxin gene. The resulting diminished expression is linked with a dying-back neuropathy impacting sensory neurons and spinocerebellar and corticospinal motor tracts (Puccio and Koenig, 2002). Frataxin is an iron chaperone required for formation of iron–sulfur (Fe–S) clusters, but its loss is associated not only with diminished activity of Fe–S-containing enzymes (important for optimal mitochondrial respiratory chain function), but also with deficient defense against oxidative stress (Bencze et al., 2006). In mouse models of the disease mitochondrial dysfunction (linked to diminished respiratory chain activity) occurs in the absence of any enhancement of oxidative stress (Seznec et al., 2005). Some clarity on the etiology of this disease has come from work in Drosophila where mitochondrial inner membrane depolarization preceded impaired mitochondrial trafficking and distal loss of fibers (Shidara and Hollenbeck, 2010). In spite of these mitochondrial abnormalities there was no sign of oxidative stress linked to respiratory chain activity. There are distinct parallels with mitochondrial dysfunction observed in diabetic neuropathy. Inner membrane depolarization in sensory neurons is identified early in the disease in animal models and respiratory chain dysfunction is observed in the absence of any attendant elevation in ROS production (Akude et al., 2011; Chowdhury et al., 2010; Huang et al., 2003; Srinivasan et al., 2000). Schwann cell mitochondria involvement in axonal degeneration in diabetes The above sections have presented an axon-centric view of mitochondrial involvement in diabetic neuropathy, however, Schwann cell abnormalities are believed to be important contributors to the etiology of nerve degeneration in diabetes (Dobrowsky et al., 2005;

61

Eckersley, 2002; Kennedy and Zochodne, 2005b). Ultrastructure studies in nerve from humans and animals reveal extensive Schwann cell degeneration combined with subtle abnormalities in mitochondria in Schwann cells (Kalichman et al., 1998; Mizisin et al., 2007). While no studies have been performed on Schwann cell mitochondrial physiology in diabetic animal models Schwann cell cultures have been exposed to high [glucose] in vitro. Cultured primary Schwann cells were treated with 30 mM glucose for 2 to 16 days and a quantitative proteomics approach was taken and revealed high [glucose]-induced increases in expression of a range of mitochondrial proteins associated with mitochondrial dysfunction, Krebs cycle, oxidative phosphorylation and detoxification; with a peak around 2–6 days (Zhang et al., 2010). Schwann cells treated with 30 mM glucose for 3 days exhibited raised rates of OCR but the efficiency of oxidative phosphorylation was impaired (Zhang et al., 2010). There was also a 1.9-fold increase in expression/activity of manganese superoxide dismutase that may explain the lack of enhanced superoxide levels. Interestingly, gene array studies on sciatic nerve from 24 week db/db diabetic mice reveal a general 2-fold enhancement in expression of 175 mitochondrially-related genes (Pande et al., 2011). This phenotype was paralleled by a 2-fold decrease in expression of 52 different mitochondrially-related mRNAs (Pande et al., 2011). The Feldman group also performed gene array screens on sural nerve from diabetic patients but in this case no significant alterations in mitochondrialrelated genes were identified (Hur et al., 2011). Caution should be exerted when interpreting this data due to the mixed cellular nature of the tissue, however, these studies provide some important insight into possible Schwann cell mitochondrial-related phenotype changes in diabetes. Glial cells within the CNS direct glucose metabolism away from oxidation through inhibition of the pyruvate dehydrogenase (PDH) complex (Halim et al., 2010). If the same regulatory restraints are present in Schwann cells then one can speculate that under high [glucose] in diabetes this inhibition may be suppressed to induce higher rates of oxidative phosphorylation, as observed by Zhang et al. (2010) — a general up-regulation in expression of mitochondrial genes would be expected to be coupled to such a process. The outcome could be excessive free radical generation, if protective mechanisms fail over time, and impairments of Schwann cell function that could translate to suboptimal axonal function. Signs of such dysfunction could be increased oxidative stress (Obrosova, 2002; Vincent et al., 2004b) and impairments in neurotrophic support (Calcutt et al., 1992; Calcutt et al., 2008; Mizisin et al., 1997). An interesting parallel to the effect of high glucose in Schwann cells is observed in rats treated with dichloroacetate (DCA), a compound that enhances activation of PDH and would be expected to raise glucose oxidation and oxidative phosphorylation in Schwann cells — although not only Schwann cells are impacted by this treatment (Stacpoole et al., 2008). Treatment with DCA caused oxidative stress in nerve but no clear effect on Schwann cell ultrastructure and had profound inhibitory effects on sensory and motor physiology and function, including significant loss of axonal caliber (Calcutt et al., 2009b). Studies from the Brady group have shown that optimal Schwann cell function is required for control of axon caliber (de Waegh et al., 1992; Kirkpatrick and Brady, 1994). Finally, a recent paper from the Milbrandt group shows elegantly that direct impairment of Schwann cell mitochondrial function leads to neuropathy in mice, but without death of Schwann cells (Viader et al., 2011). Taken together the data suggest that under normal conditions in Schwann cells a combination of aerobic and anaerobic metabolism maintains optimal ATP generation. This metabolic support is enough for Schwann cell survival (anaerobic metabolism alone can cope), to maintain nutrient delivery to the axon (e.g. transfer of lactate to the adjacent axon, as in the CNS) (Allaman et al., 2011; Vega et al., 2003), regulate myelination and provide neurotrophic support of the axon (via CNTF, for example) (Mizisin et al., 2004). Under diabetic

62

S.K.R. Chowdhury et al. / Neurobiology of Disease 51 (2013) 56–65

conditions, and in part replicated by DCA treatment, glucose oxidation is enhanced leading to a greater involvement of mitochondrial oxidative phosphorylation in ATP generation. It can be surmised that the Schwann cell is not optimized to manage high mitochondrial activity and the attendant elevation in respiratory chain-dependent superoxide production and thus there is development of oxidative stress. This stress is combined with elevated aldose reductase activity and raised polyol accumulation that also contributes to further development of oxidative stress (Ido et al., 2010; Oates, 2008). These pathological abnormalities alter Schwann cell function leading to impaired myelination (Dobrowsky et al., 2005; Kennedy and Zochodne, 2005a; Pande et al., 2011), loss of axon caliber (Jakobsen, 1976a, 1976b) and aberrant Schwann cell and axon-targeted neurotrophic support (Calcutt et al., 1992; McGuire et al., 2009; Mizisin et al., 2004; Yu et al., 2008). Until direct measures of Schwann cell mitochondrial function are attempted in the diabetic setting the above premise will remain entirely hypothetical.

Putative mechanism(s) of distal axonopathy linked to neuronal mitochondrial dysfunction The data in Fig. 2 reveals that mitochondria in diabetic sensory neurons are impaired in their ability to adapt to changes in cellular bioenergetic demand. Therefore, under conditions of high ATP demand in diabetic neurons the respiratory chain will be working at maximal capacity and our findings suggest that this workload is insufficient to comply with the energetic demands of the distal axon. The diminishment of mitochondrial inner membrane potential in the axons of diabetic neurons is expected if the flux of electron transfer through the respiratory chain is not able to cope with rates of ATP consumption (Akude et al., 2011). However, the coupling ratio was not significantly altered by the diabetic state suggesting other factors may contribute to the depolarization of the inner membrane. For example, induction of uncoupling proteins may explain the observation that proton leak was raised from 7% to 9% in diabetic neurons (Fig. 2H). This leads to the hypothesis that under such conditions depletion of the transmembrane proton gradient via pumping of protons through the ATP synthase for ATP synthesis will exceed the rate of electron transport-mediated proton translocation to the outer mitochondrial compartment. We propose that the primary cause of this reduced mitochondrial adaptability is a change in metabolic phenotype of the neuron driven by nutrient excess. Glucose uptake is insulin-independent in neurons and so a high extracellular glucose concentration will distribute equally across the plasma membrane driven via equilibrative transport mediated primarily by GLUT3 (Simpson et al., 2007; Tomlinson and Gardiner, 2008). Under diabetic conditions within the perikarya the high intracellular glucose concentration triggers the AMPK/PGC-1α pathway to be down-regulated (Akude et al., 2011; Canto and Auwerx, 2009; Chowdhury et al., 2011; Hardie, 2008). It can be postulated that this process deviates maintenance of neuronal bioenergetics away from a reliance on mitochondrial oxidative phosphorylation in favor of less energetically efficient anaerobic metabolism (glycolysis). This phenotypic change is akin to the Crabtree effect described in yeast and leads to a downregulation of respiratory chain components, matrix enzymes and mitochondrial biogenesis (Ibsen, 1961). The altered mitochondrial character is then transferred along the axon in a proximo-distal direction through anterograde transport of organelles to the nerve endings. Mitochondrial fusion of perikarya-derived newly generated or refurbished mitochondria with other more mature mitochondria along the length of the axon, combined with subsequent axonal mitochondrial fission events, will produce a homogeneous population of mitochondria with similar phenotypes (Chen and Chan, 2006). We propose that this maladaptive process causes neurodegeneration in sensory neurons, particularly those with the longest axons, because

the perikarya is unable to accurately predict the bioenergetic requirements of the distal nerve ending. Calculations of energy consumption by neurons in gray matter in the brain show that approximately 50% of ATP consumption is dedicated to action potential generation (Attwell and Laughlin, 2001). Processes such as maintenance of the resting potential, vesicular transport, Ca 2 + homeostasis and receptor cycling utilize the remainder of ATP supplies. The free nerve endings of unmyelinated and myelinated nerve fibers within the skin are very plastic and have to maintain extensive fields of innervation. Such plasticity is driven by ongoing axonal regeneration and collateral sprouting (Diamond et al., 1987; Diamond et al., 1992a; Diamond et al., 1992b) and this requires actin-treadmilling to generate force for motility of the growth cone (Bray, 1989; Bray and White, 1988). The latter process has been estimated to utilize 50% of the axons ATP reserves (Bernstein and Bamburg, 2003). There is an excellent discussion of the features of IENF plasticity by Bennett et al. in the context of distal axonopathy in chemotherapy-induced peripheral neuropathy (Bennett et al., 2011). In addition, unmyelinated axons have been found to be more energetically demanding than myelinated axons and are estimated to consume 2.5- to 10-fold more energy per action potential generated (Wang et al., 2008). Electron tomography studies on ultrastructure of mitochondria in axoplasm of peripheral nerve reveal abundant condensed mitochondria, which is indicative of high workload (e.g. high axonal demand for ATP synthesis) (Perkins and Ellisman, 2011). These concentrations of condensed mitochondria are particularly dense at the nodal region and are not observed in adjacent Schwann cells (Perkins and Ellisman, 2011). Therefore, we hypothesize that ATP consumption by the distal unmyelinated axon within the epidermis is likely to be enormous, and will fluctuate between wide thresholds and thus will outstrip the energetic requirements of the perikarya and other segments of the neuronal axis. Fig. 3 is a model that attempts to summarize the etiology of distal nerve degeneration subsequent to neuronal mitochondrial dysfunction. The consequences of suboptimal ATP supply for the distal nerve fiber are numerous: (1) collateral sprouting and plasticity will be retarded, (2) this will lead to gradual pruning of the axonal network and shrinkage of sensory innervation fields, and (3) end organs of myelinated fibers within the dermis will lose innervation and function (see Fig. 3). The mechanism(s) of axonal pruning and/or degeneration could involve a number of pathways including local activation of caspases and protein destruction (Vincent et al., 2010), impaired calcium homeostasis leading to calpain activation and protein destruction (Fernyhough and Calcutt, 2010; Gumy et al., 2010; Verkhratsky and Fernyhough, 2008), impaired Na + homeostasis (Greenwood et al., 2007), and formation of aberrant cytoskeletal structures (Bamburg et al., 2010; Bernstein et al., 2006). The latter possibility is intriguing and has been proposed to trigger formation of cytoskeletal aggregates in Alzheimer's (Bamburg et al., 2010). We suspect that this process may underlie generation of axonal swellings as seen in diabetic neuropathy (Lauria et al., 2003; Schmidt et al., 1981; Schmidt et al., 1997) and in sensory neuron cultures derived from STZ-diabetic rats (Zherebitskaya et al., 2009). Under conditions of ATP exhaustion axons of cultured embryonic hippocampal neurons form actin rods that are enriched in the dynamizing proteins actindepolymerizing factor (ADF) and cofilin (Bernstein et al., 2006). This process is usually transient and believed to serve as an in-built protective mechanism to preserve ATP pools through inhibition of actin-treadmilling and protect mitochondria from severe depolarization under high ATP demand. However, under long term exposure to suboptimal ATP levels the ADF/coflin enriched actin rods become irreversible and possibly act as loci for formation of axonal beads (or swelling) (Bamburg et al., 2010). This is an interesting and testable hypothesis that can be studied in cultures of sensory neurons derived from diabetic animals.

S.K.R. Chowdhury et al. / Neurobiology of Disease 51 (2013) 56–65

63

Fig. 3. Mitochondrial maladaption in diabetes leads to distal degeneration and loss of nerve fibers. We hypothesize that a key feature of the distal nerve fiber compared with the perikarya is a higher and more fluctuating demand for ATP. For example, collateral sprouting and axon regeneration, due to the need for growth cone motility, consume very high levels of ATP that are not paralleled in the perikarya. Due to nutrient excess in diabetes the AMPK/PGC-1α pathway is diminished and so mitochondrial phenotype throughout the sensory neuron axis is altered and becomes less capable of adapting to high peaks of ATP demand within the distal axon. Specifically, the low respiratory spare capacity and impaired respiratory control ratio means the mitochondria are poorly adapted to rapidly increase rates of electron transport under conditions requiring enhancement of the proton-motive force and associated high ATP synthase activity. This defect is highlighted by the increased state of depolarization of the mitochondrial inner membrane in axons of diabetic neurons compared with controls. The deficits in ATP generating capacity will reduce growth come motility and could trigger cytoskeletal aggregates leading to axon dystrophy (swelling) that may contribute to deficient axonal plasticity and regeneration. Within the perikarya the suboptimal ATP levels may trigger cell shrinkage and a local sprouting response that is coupled with axonal dystrophy. In chronic type 1 diabetes in rodent models over a longer time scale this cellular bioenergetic defect may trigger the irreversible loss of small unmyelinated neurons that exhibit intrinsically high ATP consumption rates. This figure is a diagrammatic representation of structural investigations of nerve fibers in humans and animal models (Jakobsen, 1976a; Lauria et al., 2003; Schmidt et al., 1997; Wendelschafer-Crabb et al., 2006). The artwork in this figure was illustrated by Mr. Daniel Mizisin, San Diego, CA, USA.

Conclusion and the future

Acknowledgments

Evidence is mounting that defects in neuronal mitochondrial function could contribute to dying-back of axons in diabetes. The distal nerve ending has unique bioenergetic requirements and under the stress of diabetic conditions the sensory neuron axis is unable to adapt and the result is neurodegenerative dissolution of the distal axon. This review has taken an axon-centric view of the mitochondrial pathways involved in diabetic neuropathy. Clearly, future work needs to be performed on mitochondrial physiology in Schwann cells in order for a more rounded view of the etiology to be forthcoming. New imaging methodologies for studying Schwann cell physiology are now available (Petrescu et al., 2007), although technically challenging, and recent studies show that mitochondrial respiration can be assessed in the nerve (Viader et al., 2011; Zheng et al., 2011). Future work is also needed in the whole animal setting to clearly delineate which components of the AMPK/PGC-1α pathway are important in diabetic neuropathy. The use of conditional mutants in sensory neurons or Schwann cells will be needed in this regard. Finally, experiments need to be designed, that will be technically demanding, to confirm that ATP exhaustion is indeed a key etiological feature in the neurodegeneration of distal fibers in diabetes.

Drs. Smith and Roy Chowdhury were supported by grants to P.F. from CIHR (grant # MOP-84214) and Juvenile Diabetes Research Foundation (grant # 1-2008-193). We thank Dwayne Morrow for technical assistance. This work was also funded by St Boniface Hospital Research. The artwork in Fig. 3 was illustrated by Mr. Daniel Mizisin, San Diego, CA, USA (email: [email protected]). References Abbate, S.L., et al., 1991. Amount and speed of fast axonal transport in diabetes. Diabetes 40, 111–117. Akude, E., et al., 2011. Diminished superoxide generation is associated with respiratory chain dysfunction and changes in the mitochondrial proteome of sensory neurons from diabetic rats. Diabetes 60, 288–297. Allaman, I., et al., 2011. Astrocyte–neuron metabolic relationships: for better and for worse. Trends Neurosci. 34, 76–87. Attwell, D., Laughlin, S.B., 2001. An energy budget for signaling in the grey matter of the brain. J. Cereb. Blood Flow Metab. 21, 1133–1145. Azzu, V., et al., 2008. High membrane potential promotes alkenal-induced mitochondrial uncoupling and influences adenine nucleotide translocase conformation. Biochem. J. 413, 323–332. Baloh, R.H., et al., 2007. Altered axonal mitochondrial transport in the pathogenesis of Charcot–Marie–Tooth disease from mitofusin 2 mutations. J. Neurosci. 27, 422–430.

64

S.K.R. Chowdhury et al. / Neurobiology of Disease 51 (2013) 56–65

Bamburg, J.R., et al., 2010. ADF/Cofilin-actin rods in neurodegenerative diseases. Curr. Alzheimer Res. 7, 241–250. Beiswenger, K.K., et al., 2008. Dissociation of thermal hypoalgesia and epidermal denervation in streptozotocin-diabetic mice. Neurosci. Lett. 442, 267–272. Bencze, K.Z., et al., 2006. The structure and function of frataxin. Crit. Rev. Biochem. Mol. Biol. 41, 269–291. Bennett, G.J., et al., 2011. Terminal arbor degeneration—a novel lesion produced by the antineoplastic agent paclitaxel. Eur. J. Neurosci. 33, 1667–1676. Bernstein, B.W., Bamburg, J.R., 2003. Actin-ATP hydrolysis is a major energy drain for neurons. J. Neurosci. 23, 1–6. Bernstein, B.W., et al., 2006. Formation of actin-ADF/cofilin rods transiently retards decline of mitochondrial potential and ATP in stressed neurons. Am. J. Physiol. Cell Physiol. 291, C828–C839. Brand, M.D., Nicholls, D.G., 2011. Assessing mitochondrial dysfunction in cells. Biochem. J. 435, 297–312. Bray, D., 1989. Growth cone formation and navigation: axonal growth. Curr. Opin. Cell Biol. 1, 87–90. Bray, D., White, J.G., 1988. Cortical flow in animal cells. Science 239, 883–888. Bugger, H., Abel, E.D., 2010. Mitochondria in the diabetic heart. Cardiovasc. Res. 88, 229–240. Bugger, H., et al., 2009. Tissue-specific remodeling of the mitochondrial proteome in type 1 diabetic Akita mice. Diabetes 58, 1986–1997. Burnand, R.C., et al., 2004. Expression of axotomy-inducible and apoptosis-related genes in sensory nerves of rats with experimental diabetes. Brain Res. Mol. Brain Res. 132, 235–240. Calcutt, N.A., et al., 1990. Axonal transport of substance P-like immunoreactivity in ganglioside-treated diabetic rats. J. Neurol. Sci. 96, 283–291. Calcutt, N.A., et al., 1992. Reduced ciliary neuronotrophic factor-like activity in nerves from diabetic or galactose-fed rats. Brain Res. 575, 320–324. Calcutt, N.A., et al., 2008. Growth factors as therapeutics for diabetic neuropathy. Curr. Drug Targets 9, 47–59. Calcutt, N.A., et al., 2009a. Therapies for hyperglycaemia-induced diabetic complications: from animal models to clinical trials. Nat. Rev. Drug Discov. 8, 417–429. Calcutt, N.A., et al., 2009b. Peripheral neuropathy in rats exposed to dichloroacetate. J. Neuropathol. Exp. Neurol. 68, 985–993. Canto, C., Auwerx, J., 2009. PGC-1alpha, SIRT1 and AMPK, an energy sensing network that controls energy expenditure. Curr. Opin. Lipidol. 20, 98–105. Chen, H., Chan, D.C., 2006. Critical dependence of neurons on mitochondrial dynamics. Curr. Opin. Cell Biol. 18, 453–459. Cheng, C., Zochodne, D.W., 2003. Sensory neurons with activated caspase-3 survive long-term experimental diabetes. Diabetes 52, 2363–2371. Chowdhury, S.K., et al., 2010. Mitochondrial respiratory chain dysfunction in dorsal root ganglia of streptozotocin-induced diabetic rats and its correction by insulin treatment. Diabetes 59, 1082–1091. Chowdhury, S.K., et al., 2011. Nutrient excess and altered mitochondrial proteome and function contribute to neurodegeneration in diabetes. Mitochondrion 11, 845–854. Christianson, J.A., et al., 2003. Restorative effects of neurotrophin treatment on diabetes-induced cutaneous axon loss in mice. Exp. Neurol. 179, 188–199. Christianson, J.A., et al., 2007. Neurotrophic modulation of myelinated cutaneous innervation and mechanical sensory loss in diabetic mice. Neuroscience 145, 303–313. de Waegh, S.M., et al., 1992. Local modulation of neurofilament phosphorylation, axonal caliber, and slow axonal transport by myelinating Schwann cells. Cell 68, 451–463. Delcroix, J.D., et al., 1998. Effect of nerve growth factor treatment on p75NTR gene expression in lumbar dorsal root ganglia of streptozocin-induced diabetic rats. Diabetes 47, 1779–1785. Diamond, J., et al., 1987. Evidence that endogenous beta nerve growth factor is responsible for the collateral sprouting, but not the regeneration, of nociceptive axons in adult rats. Proc. Natl. Acad. Sci. U. S. A. 84, 6596–6600. Diamond, J., et al., 1992a. Sensory nerves in adult rats regenerate and restore sensory function to the skin independently of endogenous NGF. J. Neurosci. 12, 1467–1476. Diamond, J., et al., 1992b. Endogenous NGF and nerve impulses regulate the collateral sprouting of sensory axons in the skin of the adult rat. J. Neurosci. 12, 1454–1466. Dobrowsky, R.T., et al., 2005. Altered neurotrophism in diabetic neuropathy: spelunking the caves of peripheral nerve. J. Pharmacol. Exp. Ther. 313, 485–491. Dominy Jr., J.E., et al., 2010. Nutrient-dependent regulation of PGC-1alpha's acetylation state and metabolic function through the enzymatic activities of Sirt1/GCN5. Biochim. Biophys. Acta 1804, 1676–1683. Ebenezer, G.J., et al., 2011. Impaired neurovascular repair in subjects with diabetes following experimental intracutaneous axotomy. Brain 134, 1853–1863. Echtay, K.S., et al., 2003. A signalling role for 4-hydroxy-2-nonenal in regulation of mitochondrial uncoupling. EMBO J. 22, 4103–4110. Eckersley, L., 2002. Role of the Schwann cell in diabetic neuropathy. Int. Rev. Neurobiol. 50, 293–321. Feige, J.N., Auwerx, J., 2007. Transcriptional coregulators in the control of energy homeostasis. Trends Cell Biol. 17, 292–301. Fernyhough, P., Calcutt, N.A., 2010. Abnormal calcium homeostasis in peripheral neuropathies. Cell Calcium 47, 130–139. Fernyhough, P., et al., 2010. Mitochondrial stress and the pathogenesis of diabetic neuropathy. Expert. Rev. Endocrinol. Metab. 5, 39–49. Francis, G., et al., 2009. Intranasal insulin ameliorates experimental diabetic neuropathy. Diabetes 58, 934–945. Greenwood, S.M., et al., 2007. Mitochondrial dysfunction and dendritic beading during neuronal toxicity. J. Biol. Chem. 282, 26235–26244.

Gumy, L.F., et al., 2010. The role of local protein synthesis and degradation in axon regeneration. Exp. Neurol. 223, 28–37. Hahn, K., et al., 2008. Differential effects of HIV infected macrophages on dorsal root ganglia neurons and axons. Exp. Neurol. 210, 30–40. Halim, N.D., et al., 2010. Phosphorylation status of pyruvate dehydrogenase distinguishes metabolic phenotypes of cultured rat brain astrocytes and neurons. Glia 58, 1168–1176. Hardie, D.G., 2008. AMPK: a key regulator of energy balance in the single cell and the whole organism. Int. J. Obes. (Lond) 32 (Suppl. 4), S7–S12. Hellweg, R., et al., 1994. Axonal transport of endogenous nerve growth factor (NGF) and NGF receptor in experimental diabetic neuropathy. Exp. Neurol. 130, 24–30. Hill, B.G., et al., 2009. Importance of the bioenergetic reserve capacity in response to cardiomyocyte stress induced by 4-hydroxynonenal. Biochem. J. 424, 99–107. Huang, T.J., et al., 2003. Insulin prevents depolarization of the mitochondrial inner membrane in sensory neurons of type 1 diabetic rats in the presence of sustained hyperglycemia. Diabetes 52, 2129–2136. Huang, T.J., et al., 2005a. Neurotrophin-3 prevents mitochondrial dysfunction in sensory neurons of streptozotocin-diabetic rats. Exp. Neurol. 194, 279–283. Huang, T.J., et al., 2005b. Insulin enhances mitochondrial inner membrane potential and increases ATP levels through phosphoinositide 3-kinase in adult sensory neurons. Mol. Cell. Neurosci. 28, 42–54. Hur, J., et al., 2011. The identification of gene expression profiles associated with progression of human diabetic neuropathy. Brain 134, 3222–3235. Ibsen, H.K., 1961. The Crabtree effect: a review. Cancer Res. 21, 829–841. Ido, Y., et al., 2010. Early neural and vascular dysfunctions in diabetic rats are largely sequelae of increased sorbitol oxidation. Antioxid. Redox Signal. 12, 39–51. Jakobsen, J., 1976a. Axonal dwindling in early experimental diabetes. I. A study of cross sectioned nerves. Diabetologia 12, 539–546. Jakobsen, J., 1976b. Axonal dwindling in early experimental diabetes. II. A study of isolated nerve fibres. Diabetologia 12, 547–553. Jekabsons, M.B., Nicholls, D.G., 2004. In situ respiration and bioenergetic status of mitochondria in primary cerebellar granule neuronal cultures exposed continuously to glutamate. J. Biol. Chem. 279, 32989–33000. Jiang, Y., et al., 2004. Selective loss of calcitonin gene-related Peptide-expressing primary sensory neurons of the a-cell phenotype in early experimental diabetes. Diabetes 53, 2669–2675. Jolivalt, C.G., et al., 2008. Allodynia and hyperalgesia in diabetic rats are mediated by GABA and depletion of spinal potassium-chloride co-transporters. Pain 140, 48–57. Kalichman, M.W., et al., 1998. Reactive, degenerative, and proliferative Schwann cell responses in experimental galactose and human diabetic neuropathy. Acta Neuropathol. 95, 47–56. Kamiya, H., et al., 2006. Degeneration of the Golgi and neuronal loss in dorsal root ganglia in diabetic BioBreeding/Worcester rats. Diabetologia 49, 2763–2774. Kennedy, J.M., Zochodne, D.W., 2005a. Experimental diabetic neuropathy with spontaneous recovery: is there irreparable damage? Diabetes 54, 830–837. Kennedy, J.M., Zochodne, D.W., 2005b. Impaired peripheral nerve regeneration in diabetes mellitus. J. Peripher. Nerv. Syst. 10, 144–157. Kennedy, W.R., et al., 1996. Quantitation of epidermal nerves in diabetic neuropathy. Neurology 47, 1042–1048. Kilgour, R.D., et al., 1990. Diabetes affects retrograde but not anterograde transport of sciatic nerve phosphofructokinase in Sprague–Dawley rats. Can. J. Physiol. Pharmacol. 68, 1317–1321. Kirkpatrick, L.L., Brady, S.T., 1994. Modulation of the axonal microtubule cytoskeleton by myelinating Schwann cells. J. Neurosci. 14, 7440–7450. Lauria, G., et al., 2003. Axonal swellings predict the degeneration of epidermal nerve fibers in painful neuropathies. Neurology 61, 631–636. Lehmann, H.C., et al., 2011. Mitochondrial dysfunction in distal axons contributes to human immunodeficiency virus sensory neuropathy. Ann. Neurol. 69, 100–110. Malik, R.A., et al., 2005. Sural nerve pathology in diabetic patients with minimal but progressive neuropathy. Diabetologia 48, 578–585. McGuire, J.F., et al., 2009. Caveolin-1 and altered neuregulin signaling contribute to the pathophysiological progression of diabetic peripheral neuropathy. Diabetes 58, 2677–2686. Misko, A., et al., 2010. Mitofusin 2 is necessary for transport of axonal mitochondria and interacts with the Miro/Milton complex. J. Neurosci. 30, 4232–4240. Mizisin, A.P., et al., 1997. Aldose reductase inhibition increases CNTF-like bioactivity and protein in sciatic nerves from galactose-fed and normal rats. Diabetes 46, 647–652. Mizisin, A.P., et al., 2004. Ciliary neurotrophic factor improves nerve conduction and ameliorates regeneration deficits in diabetic rats. Diabetes 53, 1807–1812. Mizisin, A.P., et al., 2007. Comparable myelinated nerve pathology in feline and human diabetes mellitus. Acta Neuropathol. 113, 431–442. Mootha, V.K., et al., 2003. PGC-1alpha-responsive genes involved in oxidative phosphorylation are coordinately downregulated in human diabetes. Nat. Genet. 34, 267–273. Murphy, M.P., et al., 2003. Superoxide activates uncoupling proteins by generating carbon-centered radicals and initiating lipid peroxidation: studies using a mitochondria-targeted spin trap derived from alpha-phenyl-N-tert-butylnitrone. J. Biol. Chem. 278, 48534–48545. Nicholls, D.G., 2006. Simultaneous monitoring of ionophore- and inhibitor-mediated plasma and mitochondrial membrane potential changes in cultured neurons. J. Biol. Chem. 281, 14864–14874. Nja, A., Purves, D., 1978. The effects of nerve growth factor and its antiserum on synapses in the superior cervical ganglion of the guinea-pig. J. Physiol. 277, 53–75. Oates, P.J., 2008. Aldose reductase, still a compelling target for diabetic neuropathy. Curr. Drug Targets 9, 14–36.

S.K.R. Chowdhury et al. / Neurobiology of Disease 51 (2013) 56–65 Obrosova, I.G., 2002. How does glucose generate oxidative stress in peripheral nerve? Int. Rev. Neurobiol. 50, 3–35. Pande, M., et al., 2011. Transcriptional profiling of diabetic neuropathy in the BKS db/db mouse: a model of type 2 diabetes. Diabetes 60, 1981–1989. Patti, M.E., et al., 2003. Coordinated reduction of genes of oxidative metabolism in humans with insulin resistance and diabetes: potential role of PGC1 and NRF1. Proc. Natl. Acad. Sci. U. S. A. 100, 8466–8471. Perkins, G.A., Ellisman, M.H., 2011. Mitochondrial configurations in peripheral nerve suggest differential ATP production. J. Struct. Biol. 173, 117–127. Petrescu, N., et al., 2007. Sources of axonal calcium loading during in vitro ischemia of rat dorsal roots. Muscle Nerve 35, 451–457. Polydefkis, M., et al., 2004. The time course of epidermal nerve fibre regeneration: studies in normal controls and in people with diabetes, with and without neuropathy. Brain 127, 1606–1615. Puccio, H., Koenig, M., 2002. Friedreich ataxia: a paradigm for mitochondrial diseases. Curr. Opin. Genet. Dev. 12, 272–277. Puigserver, P., et al., 1998. A cold-inducible coactivator of nuclear receptors linked to adaptive thermogenesis. Cell 92, 829–839. Quattrini, C., et al., 2007. Surrogate markers of small fiber damage in human diabetic neuropathy. Diabetes 56, 2148–2154. Richardson, D.K., et al., 2005. Lipid infusion decreases the expression of nuclear encoded mitochondrial genes and increases the expression of extracellular matrix genes in human skeletal muscle. J. Biol. Chem. 280, 10290–10297. Rodgers, J.T., et al., 2008. Metabolic adaptations through the PGC-1 alpha and SIRT1 pathways. FEBS Lett. 582, 46–53. Roy Chowdhury, S.K., Smith, D.R., Saleh, A., Schapansky, J., Gomes, S., Akude, E., Morrow, D., Calcutt, N.A., Fernyhough, P., in press. Impaired AMP-activated protein kinase signaling in dorsal root ganglia neurons is linked to mitochondrial dysfunction and peripheral neuropathy in diabetes. Brain. Said, G., 2007. Diabetic neuropathy—a review. Nat. Clin. Pract. Neurol. 3, 331–340. Schmidt, R.E., 2002. Neuropathology and pathogenesis of diabetic autonomic neuropathy. Int. Rev. Neurobiol. 50, 257–292. Schmidt, R.E., et al., 1981. Experimental diabetic autonomic neuropathy. Am. J. Pathol. 103, 210–225. Schmidt, R.E., et al., 1987. Orthograde and retrograde axonal transport of dopaminebeta-hydroxylase in ileal mesenteric nerves of rats with chronic streptozotocin diabetes. Brain Res. 401, 142–146. Schmidt, R.E., et al., 1997. Dystrophic axonal swellings develop as a function of age and diabetes in human dorsal root ganglia. J. Neuropathol. Exp. Neurol. 56, 1028–1043. Schmidt, R.E., et al., 2003. Non-obese diabetic mice rapidly develop dramatic sympathetic neuritic dystrophy: a new experimental model of diabetic autonomic neuropathy. Am. J. Pathol. 163, 2077–2091. Schmidt, R.E., et al., 2008. Synaptic ultrastructural alterations anticipate the development of neuroaxonal dystrophy in sympathetic ganglia of aged and diabetic mice. J. Neuropathol. Exp. Neurol. 67, 1166–1186. Schmidt, R.E., et al., 2009. Neuritic dystrophy and neuronopathy in Akita (Ins2(Akita)) diabetic mouse sympathetic ganglia. Exp. Neurol. 216, 207–218. Schmidt, R.E., et al., 2011. Effect of insulin and an erythropoietin-derived peptide (ARA290) on established neuritic dystrophy and neuronopathy in Akita (Ins2 Akita) diabetic mouse sympathetic ganglia. Exp. Neurol. 232, 126–135. Seznec, H., et al., 2005. Friedreich ataxia: the oxidative stress paradox. Hum. Mol. Genet. 14, 463–474. Shidara, Y., Hollenbeck, P.J., 2010. Defects in mitochondrial axonal transport and membrane potential without increased reactive oxygen species production in a Drosophila model of Friedreich ataxia. J. Neurosci. 30, 11369–11378.

65

Sidenius, P., Jakobsen, J., 1980. Reduced perikaryal volume of lower motor and primary sensory neurons in early experimental diabetes. Diabetes 29, 182–186. Simpson, I.A., et al., 2007. Supply and demand in cerebral energy metabolism: the role of nutrient transporters. J. Cereb. Blood Flow Metab. 27, 1766–1791. Sivitz, W.I., Yorek, M.A., 2010. Mitochondrial dysfunction in diabetes: from molecular mechanisms to functional significance and therapeutic opportunities. Antioxid. Redox Signal. 12, 537–577. Srinivasan, S., et al., 2000. Diabetic peripheral neuropathy: evidence for apoptosis and associated mitochondrial dysfunction. Diabetes 49, 1932–1938. Stacpoole, P.W., et al., 2008. Role of dichloroacetate in the treatment of genetic mitochondrial diseases. Adv. Drug Deliv. Rev. 60, 1478–1487. Sugimoto, K., et al., 2008. Role of advanced glycation end products in diabetic neuropathy. Curr. Pharm. Des. 14, 953–961. Szendroedi, J., et al., 2011. The role of mitochondria in insulin resistance and type 2 diabetes mellitus. Nat. Rev. Endocrinol. 8, 92–103. Tomlinson, D.R., Gardiner, N.J., 2008. Glucose neurotoxicity. Nat. Rev. Neurosci. 9, 36–45. Vega, C., et al., 2003. Uptake of locally applied deoxyglucose, glucose and lactate by axons and Schwann cells of rat vagus nerve. J. Physiol. 546, 551–564. Verkhratsky, A., Fernyhough, P., 2008. Mitochondrial malfunction and Ca2 + dyshomeostasis drive neuronal pathology in diabetes. Cell Calcium 44, 112–122. Viader, A., et al., 2011. Schwann cell mitochondrial metabolism supports long-term axonal survival and peripheral nerve function. J. Neurosci. 31, 10128–10140. Vincent, A.M., et al., 2004a. Uncoupling proteins prevent glucose-induced neuronal oxidative stress and programmed cell death. Diabetes 53, 726–734. Vincent, A.M., et al., 2004b. Oxidative stress in the pathogenesis of diabetic neuropathy. Endocr. Rev. 25, 612–628. Vincent, A.M., et al., 2009. Hyperlipidemia: a new therapeutic target for diabetic neuropathy. J. Peripher. Nerv. Syst. 14, 257–267. Vincent, A.M., et al., 2010. Mitochondrial biogenesis and fission in axons in cell culture and animal models of diabetic neuropathy. Acta Neuropathol. 120, 477–489. Wang, S.S., et al., 2008. Functional trade-offs in white matter axonal scaling. J. Neurosci. 28, 4047–4056. Wendelschafer-Crabb, G., et al., 2006. Morphological features of nerves in skin biopsies. J. Neurol. Sci. 242, 15–21. Xiao, W.H., et al., 2011. Mitochondrial abnormality in sensory, but not motor, axons in paclitaxel-evoked painful peripheral neuropathy in the rat. Neuroscience 199, 461–469. Yagihashi, S., 1996. Pathology and pathogenetic mechanisms of diabetic neuropathy. Diabetes Metab. Rev. 11, 193–225. Yu, C., et al., 2008. Hyperglycemia and downregulation of caveolin-1 enhance neuregulin-induced demyelination. Glia 56, 877–887. Zhang, L., et al., 2010. Hyperglycemia alters the Schwann cell mitochondrial proteome and decreases coupled respiration in the absence of superoxide production. J. Proteome Res. 9, 458–471. Zheng, H., et al., 2011. Functional deficits in peripheral nerve mitochondria in rats with paclitaxel- and oxaliplatin-evoked painful peripheral neuropathy. Exp. Neurol. 232, 154–161. Zherebitskaya, E., et al., 2009. Development of selective axonopathy in adult sensory neurons isolated from diabetic rats: role of glucose-induced oxidative stress. Diabetes 58, 1356–1364. Zuchner, S., et al., 2004. Mutations in the mitochondrial GTPase mitofusin 2 cause Charcot–Marie–Tooth neuropathy type 2A. Nat. Genet. 36, 449–451.