Cholinergic Axonal Dystrophy and Mitochondrial Pathology in Prosimian Primates

EXPERIMENTAL NEUROLOGY ARTICLE NO.

142, 111–127 (1996)

0183

Cholinergic Axonal Dystrophy and Mitochondrial Pathology in Prosimian Primates DONALD E. SCHMECHEL,1 D. SCOTT BURKHART, RUBY ANGE,

AND

M. KAY IZARD

Department of Medicine (Neurology) and Department of Neurobiology, Duke University Medical Center, Durham, North Carolina 22710; and Durham VA Medical Center, Durham, North Carolina 22710

INTRODUCTION Progressive cholinergic axonal dystrophy, cholinergic denervation, and generalized gliosis begin in the prosimian primate species Otolemur at 10% of maximum life span. In these same animals, extensive cerebral b-amyloidosis follows relatively more abruptly at 50% of maximum life span. In contrast, even at maximum life span, the prosimian primate species Galago senegalensis Moholi, Microcebus murinus, and Eulemur fulvus collaris and insectivore species T. belangeri are either spared or much less affected. In this report, we further document this progressive cholinergic denervation in Otolemur which involves first projections of the pedunculopontine nucleus (PPN, CH5-6) and later projections of CH1-4 cholinergic nuclei, as well as other noncholinergic pathways. Affected cholinergic cell bodies and axons contain abnormal mitochondria with increased content of manganese superoxide dismutase (MnSOD). This syndrome correlates with moderate copper deficiency marked by diminished liver copper levels and cuproenzyme activities, carnitine deficiency possibly secondary to renal Fanconi syndrome, and evidence for stress inflammatory response activation. Mitochondrial pathology was observed in pancreatic islet cells, proximal renal tubule epithelial cells, and choroid plexus epithelial cells, and it involved central cholinergic neurons. In Otolemur garnetti, the degree of central cholinergic injury directly correlated to depression of liver copper stores. The Otolemur syndrome involves ‘‘sentinel’’ central cholinergic injury and selective mitochondrial pathology in cell classes defined by high mitochondrial content and/or metabolic activity and high content of nitric oxide synthetase and MnSOD. Environmental factors affecting copper and carnitine metabolism could interact with genetic defects or traits to produce abnormal and aggressive aging of Otolemur. Subclinical, cellclass specific mitochondrial dysfunction in these prosimian primates may be a model for human neurodegenerative diseases. r 1996 Academic Press, Inc.

1 To whom correspondence should be addressed. Fax: (919) 6846514.

The prosimian primate species Otolemur garnetti and Otolemur crassicaudatus show aggressive brain aging beginning in early adulthood (34). This syndrome includes onset of dystrophic changes in axons of the cholinergic neurons of the midbrain pedunculopontine nucleus (PPN, CH5-6) and significant gliosis with ferritin positive microglial reaction, MHC class II activation, and astrogliosis throughout the neuraxis beginning at 10% of maximum life span (34). The dystrophic cholinergic axons are particularly prominent in the dense projection of PPN to the nucleus reticularis thalami (nRT). The extent of cholinergic dystrophy and gliosis is steadily progressive and these same species also consistently show the relatively sudden onset of significant b-amyloidosis with plaques and vascular amyloid at 50% of maximum life span. Concurrent cholinergic injury, gliosis, and b-amyloidosis in Otolemur offer a partial animal model for Alzheimer’s disease and related neurodegenerative disorders (34). This aggressive syndrome of brain aging in Otolemur does not apparently represent normal brain aging, since other prosimian species such as Galago senegalensis Moholi, Microcebus murinus, and Eulemur fulvus collaris and the related insectivore species Tupaia belangeri show much less cholinergic dystrophy, show a slower rate of gliosis, and have either no or minimal b-amyloidosis at the end of maximal life span (34). In this report, we characterize further the nature of the cholinergic injury in Otolemur particularly the prominence of mitochondrial structural abnormalities and increased content of mitochondrial superoxide dismutase (MnSOD) visible in affected cell bodies and axons. The occurrence of mitochondrial pathology even in young animals led to a systematic anatomical investigation of mitochondria in other tissues and organs. Because of the well-described syndrome of increased iron stores in prosimian species (1, 15) and the concern that reactive oxygen species (ROS) might contribute to the Otolemur syndrome, we gathered information on the status of copper metabolism, free radical protective enzymes, and lipid status in these same species.

111

0014-4886/96 $18.00 Copyright r 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.

112

SCHMECHEL ET AL.

The results support the hypothesis that aggressive brain aging in Otolemurs and prosimian primate species is closely related in age of onset, extent, and cell classes affected to mitochondrial pathology and abnormal copper and carnitine metabolism. Affected cell classes in the brain and periphery share the characteristics of high mitochondrial content and/or metabolic rate, high nitric oxide synthase content, and increased MnSOD content. Our data support the hypothesis that low tissue copper stores and (or) carnitine deficiency are closely related to the severity of this syndrome in a given species and to the observed species differences. We will discuss the possibility that genetic traits and/or environmental factors might be causally related to this syndrome, whether this may represent a mitochondrial disorder, and the significance for other animal models and for human neurodegenerative disorders. METHODS

Animals E. fulvus collaris and M. murinus tissues were collected over a 14-year period from very old animals at DUPC electively euthanized by veterinarians for health reasons of age-related decline or serious illness. Most specimens of nonendangered species (Galago, Otolemur, and Tupaia species) were electively euthanized for anatomical studies in the Department of Psychology or Department of Neurobiology, although some animals were euthanized for reasons of age-related decline and/or serious illness. The collected specimens included three wild-caught and 16 colony bred G. senegalensis Moholi (lesser bushbaby), 11 wild-caught and 49 colony bred O. garnetti (brown or greater bushbaby), two wild-caught and 53 colony bred Otolemur crassicaudatus monteiri and 23 colony bred Otolemur crassicaudatus argentatus (thick-tailed bushbaby), six M. murinus (lesser mouse lemur) aged 10–12 years old, euthanized for poor health, and three E. fulvus collaris (collared brown lemur) aged 15, 20, and 35 years of age, euthanized for poor health. These animals were kept in AALAC and/or USDA approved facilities with the nonendangered Galagidae species and Tupaia kept in breeding colonies at Duke University Medical Center and with the two lemur species kept at Duke University Primate Center, Duke University (1).

fitness and maintained hematocrit, but do run elevated serum iron (200–300 µg/dliter) and iron saturation values (50–60%) compared to other primate species (1, 15). Tissue Collection Animals were gently restrained, administered anesthesia, and then euthanized under approved protocols and veterinary supervision by lethal deep anesthesia and blood samples were taken from cardiac puncture after death. Some blood samples were taken from healthy adults under gentle restraint through femoral venipuncture. All organs were dissected and weighed, and portions were separated for immersion fixation in 10% buffered formalin and for rapid freezing in liquid nitrogen. Fixed tissue was immersed for 1–2 days and then stored in buffer with 0.1% formalin. A number of the nonendangered animals were utilized in anatomical experiments and were prepared by perfusion fixation with 10% buffered formalin (2–4 cc/g) followed after 30 min to 1 h by 10% buffered sucrose for cryoprotection and serial sectioning. Selected animals (nonendangered species) for immunoelectronmicroscopy were perfused with 4% paraformaldehyde with 0.1–0.2% EM-grade glutaraldehyde (4–6 cc/g) followed by dissection at 2–4 h and overnight fixation in thinner tissue slabs. Routine Electron Microscopy Small blocks were dissected from liver, body of pancreas, heart (left ventricle), kidney (cortex adjacent to capsule), spleen, thymus, and brain (choroid plexus) and submitted for routine imbedding and processing for transmission electron microscopy. All tissues except for liver showed reasonable ultrastructural morphology after immersion fixation described above. Midbrain and thalamus in 2 Otolemur were analyzed by flat imbedding of thicker vibratome sections (see Immunocytochemistry) to allow precise dissection of regions of PPN-LDT, oculomotor nucleus, and nRT. These dissected areas were then remounted on blank capsules for sectioning. Standard sections were collected on nickel grids and examined on a JEOL 1200EX2 electron microscope. Ultrastructural analysis included kidney tissue from 15 O. crassicaudatus, 3 O. garnetti, 4 tree shrew, and 2 M. murinus. Pancreas and choroid plexus was examined in 8 O. crassicaudatus.

Diets Diets were based on monkey food pellets, typically New World Monkey Chow or Monkey Chow (Purina), supplemented by fruit and insects (Galago, Otolemurs, and Microcebus). The iron content of these food pellets approximates 300–500 ppm of iron (1, 15). Food and water were presented ad libitum. Prosimians on these diets are generally healthy with adequate reproductive

Histology, Histochemistry, and Immunocytochemistry Oil red O staining and Perl’s reaction for iron were performed on vibratome sections of liver and kidney. Routine histological stains (hematoxylin and eosin, Mallory trichrome, periodic acid–Schiff stain, aldehydefuchsin) were performed on representative paraffin sections of liver, muscle, heart, kidney, pancreas, and

CHOLINERGIC AXONAL DYSTROPHY IN PROSIMIANS

brain. Diaminobenzidine enhanced Perl’s stain was used for liver and brain tissue to detect tissue iron (26). NADPH-diaphorase staining was performed on vibratome sections of pancreas, kidney, and brain and cryostat sections of choroid plexus (10, 42). Immunocytochemistry was performed on floating vibratome sections (25–40 µm) or freezing microtome sections (40–60 µm) for localization of choline acetyl transferase (ChAT) (rat monoclonal anti-human ChAT, Boehringer-Mannheim), serotonin (rabbit anti-5HT, Immunonuclear), tyrosine hydroxylase (rabbit anti-rat TH, Eugene Tech), glutamic acid decarboxylase (sheep anti-rat GAD) (27), manganese superoxide dismutase (rabbit anti-rat MnSOD, courtesy of Dr. L. Y. Chang, DUMC) (7, 39), ferritin (rabbit anti-human ferritin, Boehringer-Mannheim), and other markers (34). Detection methods used standard avidin–biotin kits (Vector Laboratories, Burlingame, CA) and for ChAT and MnSOD utilized enhancement with nickel–cobalt for some runs (27). Double antigen localization employed successive runs for ChAT (diaminobenzidine brown first reaction) and MnSOD (o-dianisidine green, second reaction) with method controls for each step (13). Immunoelectron Microscopy Sixty- to eighty-micrometer vibratome sections from perfused animals were immunoreacted for localization of ChAT using diaminobenzidine as chromogen, osmicated, and then flat-imbedded under Mylar plastic. Selected regions were selected, cut out and mounted on blank capsules for sectioning. Thin sections were unstained or lightly stained to allow maximal detection of electron-dense ChAT reaction product. Histological Analysis Analysis of immunocytochemical results for dystrophic and normal ChAT terminals was carried out by drawing selected fields using camera lucida using a 40X objective (dystrophic fibers) or a 100X objective with oil immersion (normal fibers). Features were drawn onto gridded paper representing 160 grid squares covering roughly 0.25 mm2 (40X objective). Three independent fields for each region for each animal were drawn and average density (squares with feature/total squares) was reported. Variation for nRT of same animal was usually 10–15% and interobserver variability tested for four animals was roughly 10% (34). L-Carnitine

Analysis

Plasma carnitine levels were determined at the Duke Mass Spectrophotometry Facility using regular clinical protocols yielding total carnitine and free and shortchain acylcarnitine levels (nM/ml). Several plasma acylcarnitine profiles were also run using internal deuterated standards to assay for the presence of

113

longer chain acylcarnitines. Tissue carnitine levels were measured on 100- to 200-mg samples submitted frozen to Metabolic Analysis Laboratories, Inc. (Madison, WI) for measurement using radioenzymatic assay with carnitine acetyltransferase to yield free carnitine and total carnitine by analysis after hydrolysis of acylcarnitines, and indirectly to yield acylcarnitine (28). Tissue carnitine levels are expressed as nanomoles per milligram noncollagen protein (NCP). Liver Metal Determinations Frozen liver samples (0.5–1.0 g) were analyzed for metal content by three methods: (i) neutron activation analysis of whole tissue samples (Nuclear Reactor Service, North Carolina State University), (ii) analysis of wet-ashed samples (3 N HCl–10% trichloroacetic acid) by atomic absorption spectroscopy using inductively coupled argon furnace (Jarrell-Ash AtomComp 1100, Compuchem, RTP, NC), and (iii) analysis of wet ashed samples (successive 10% HCl–10% HNO3) by inductively coupled argon plasma mass spectrophotometry (VG Elemental Model PQ1, Coors Analytical Laboratories). Standards included in each run consisted of NBS bovine liver standard NBS 1577a (National Bureau of Standards, Gaithersburg, MD), as well as method controls and blanks and internally spiked samples. Values are expressed as micrograms per gram wet weight (parts per million). Reference values for NBS standard are 194 6 20 ppm iron, 158 6 7 ppm copper, and 123 6 8 ppm zinc. Experimentally observed values differed by less than 5% from the above values. Liver Cu,Zn-SOD, MnSOD, and Cytochrome Oxidase Activity Frozen liver tissue samples (approximately 100 mg) were thawed, weighed, and gently homogenized in 10 volumes of 0.32 M ice-cold sucrose using microsonication (approximately 1 min) and centrifuged at 12,000g for 5 min and aliquots were taken for hemoglobin, protein, and enzyme determinations. SOD enzyme activity was determined by inhibition of ferricytochrome c reduction by superoxide produced by xanthine oxidase (22). The high and low KCN method was used to distinguish between total SOD activity (low KCN, 10 µM ) and MnSOD activity (high KCN, 1 mM with 90% Cu,Zn-SOD inhibition). Cu,Zn-SOD activity is obtained by subtraction. Protein assays were done with a kit (Pierce Biochemicals), and activities were expressed as units per milligram wet weight. Cytochrome oxidase activity was performed on liver homogenates using reduced cytochrome c (reduced by dithionite and purified on Sephadex G-25 column) as substrate by monitoring the decreased absorbance of cytochrome c during oxidation. Activity was expressed as micromoles per minute per milligram protein (3, 30, 40).

114

SCHMECHEL ET AL.

Serum Metal, Ceruloplasmin, and Vitamin E Levels Blood was collected by venipuncture in metal-free vacutainer tubes, allowed to clot, and immediately separated into serum which was flash-frozen in liquid nitrogen for storage. Determination of serum copper and zinc was carried out by flame atomic spectrophotometry and expressed as micrograms per deciliter and immunoassay of serum ceruloplasmin was carried out by turbidimetry in Duke Clinical Laboratories. Serum ceruloplasmin enzyme activity was determined by odianisidine activity (36) and by ferroxidase II activity (11) and expressed as units per liter. Serum vitamin E levels were performed by Roche Laboratories. Liver L-Ascorbic Acid Levels Frozen liver tissue samples were thawed, weighed, and gently homogenized with a glass-Teflon homogenizer in mobile phase buffer without TBA. L-Ascorbic acid levels were determined by the reverse-phase HPLC method with electrochemical detection using a mobile phase of 8 mM Na2HPO4–1 mM EDTA–5% methanol– 700 µl/liter tributylamine (TBA) using two injections for each of two digests per animal. Analysis was by isocratic separation on Alltech Hypersil BDS column C-18 3U (150X4.6, No. 1178) with a guard cell, using a Perkin–Elmer BIO 410 LC pump and an ESA Coulchem II Electrochemical Detector. Standards were freshly prepared and spectrophotometrically analyzed from L-ascorbic acid, and experimental values were determined by peak integration with a Perkin–Elmer Nelson 1020 Integrator and expressed as micrograms per gram wet weight. Data Storage and Statistical Analysis All demographic and numerical data on animals used in this study were collated and analyzed using a Statgraphics package using multiple analysis of variance and linear regressions. RESULTS

Abnormalities of Trace Mineral and Carnitine Metabolism in Prosimian Primates A common finding in all prosimian species was increased iron stores in liver and spleen as reported previously for lemur species (15). This increase is apparently associated with excess absorption from iron-enriched dietary sources common in captivity (1, 15, see Diets). Duodenal tissue from these animals is often slightly tan colored and is strongly Perl’s stain positive for iron (Fig. 1A). Tips of duodenal villi contain strongly ferritin and Prussian blue positive macrophages, supportive of excess dietary iron absorption. Liver tissue in these species displays strong Perl’s

staining for iron (Fig. 1B), and is strongly stained for fat by oil red O stains (Fig. 1C) compared to negative staining of liver of common laboratory rodent species or captive Old World primates (data not shown). The histological findings of increased liver iron stores and hepatic steatosis in these species led to the decision to obtain tissue determinations of iron, copper, zinc, and carnitine levels and to examine various body organs and tissues for pathology (e.g., pancreas relevant to iron overload (15), kidney and muscle relevant to carnitine status (17, 32, 41)). Liver iron is elevated in all prosimian species compared to captive Old World primates (usual values 200–300 µg/g wet wt), except for G. senegalensis (Table 1). Liver copper levels are significantly lower (P , 0.0001) in G. senegalensis and Otolemur species compared to Tupaia, lemur (Table 1), and Old World primate species (usual values 7–10 µg/g wet weight). Liver zinc values and zinc/copper ratio were increased in all prosimian species, particularly endangered lemur species euthanized for poor health. Determinations were also made of liver carnitine levels since hepatic steatosis suggested a possible disorder of fatty acid metabolism (Table 1) and is a feature of carnitine deficiency (24). Liver carnitine levels were significantly low in Otolemur species, intermediate in G. senegalensis, and comparable in Tupaia and the two lemur species to levels observed in laboratory rodents and other primate species (1–2 nM/mg NCP) (P , 0.0001). In Otolemur, plasma acylcarnitine profiles were normal with dominant acylcarnitine being acetylcarnitine. Urine screens showed no evidence of organic aciduria. Plasma total carnitine values were correspondingly low (usually 10–20 nM/ml) in Otolemur and correlated closely with liver carnitine. Skeletal muscle carnitine values were also low (2–10 nM/mg NCP), but correlated less well with plasma values. As is true for liver carnitine levels (Table 1), Otolemur crassicaudatus had significantly lower plasma and muscle carnitine levels than O. garnetti. Pilot experiments have demonstrated that brief oral L-carnitine supplementation (100 mg/kg for 2 weeks) can boost plasma and tissue carnitine levels in Otolemur for periods of several months. In terms of hepatic function, comparison of the two Otolemur species demonstrates relative nonketotic hypoglycemia in O. crassicaudatus after fasting greater than 24 h without apparent encephalopathy or myopathy (data not shown). In both Otolemur species, serum triglyceride levels vary inversely with liver carnitine levels (data not shown). Thus, depressed carnitine status in Otolemurs appears to be physiologically significant, but subclinical in terms of health effects. Since ascorbate deficiency can influence carnitine status (1, 32, 33), liver ascorbate levels were determined. Liver ascorbate varied significantly (Table 1), but the lowest values were not observed in Otolemur

CHOLINERGIC AXONAL DYSTROPHY IN PROSIMIANS

115

FIG. 1. (A) Strongly blue positive Perl’s stain of whole duodenal mucosa from O. garnetti (left) compared to normal pale appearance of gastric mucosa (right). (B) Perl’s stain of liver section from M. murinus indicating excess iron stores. (C) Oil red O staining of liver section from M. murinus indicating steatosis. (D) Perl’s stain with DAB enhancement showing prominent iron staining in white matter and perforant pathway (pp) of hippocampus (hc) of adult Otolemur. (E) NADPH-diaphorase histochemistry of kidney cortex from M. murinus showing minimal NOS activity. (F) Young adult Otolemur (1 year old) showing strongly positive diaphorase activity in proximal renal tubules. (G) Older adult Otolemur (11 years old) showing strongly positive diaphorase activity of tubular epithelium and possible dystrophic nerve fiber indicated by arrow. (H) Perl’s stain with DAB enhancement of frontal section of thalamus from Otolemur showing strong staining for iron in nRT and less staining in medial thalamus (thal). (I–K) Combined immunolocalization of ChAT (brown) and MnSOD (green) in Otolemur brain stem (yellow background is nonspecific chromogen staining). (I) Green stippled MnSOD staining of mitochondria in cholinergic neurons of oculomotor nucleus with omission of ChAT antibody. (J) Semiadjacent section of oculomotor nucleus showing effect of ChAT antibody and combined immunolocalization of MnSOD and ChAT. (K) Markedly increased dark green MnSOD staining of specifically identified light brown cholinergic neurons of pedunculopontine complex. (L) Similar combined immunolocalization of MnSOD and ChAT in section of basal forebrain of Otolemur showing ‘‘normal’’ appearing light green mitochondrial staining in large doubly stained cholinergic neuron (bottom center) and dark green immunoreactivity of larger, intensely stained mitochondrial profiles present in adjacent dystrophic light brown cholinergic processes including one superimposed on cell. Original magnifications: (B, C, E, F, G, I, J, K) 803; (D, H) 403; (L) 2503.

species with low liver copper or carnitine, but rather in Microcebus with high liver iron levels. The low liver copper values of 3–5 µg/g wet weight in Galago and Otolemur species do not by themselves indicate copper deficiency and are comparable to many ‘‘normal’’ rodent species (30). We therefore measured serum copper, zinc levels, the serum cuproenzyme/ copper transport protein ceruloplasmin (both usual diamine oxidase and ferroxidase II activity), and the liver cuproenzymes cytochrome oxidase and Cu,ZnSOD in these same animals (Table 1). Severe copper deficiency in rodents is accompanied by virtually absent ceruloplasmin activity, depressed serum copper, and low liver cytochrome oxidase and Cu,Zn-SOD activities (30). In fact, Otolemurs and G. senegalensis had low ceruloplasmin levels as measured by diamine oxidase activity compared to other prosimian and primate species (Table 1). Likewise, liver cytochrome oxidase and Cu,Zn-SOD enzyme activities in O. crassicaudatus and O. garnetti were low compared to those in

G. senegalensis and M. murinus. However, Otolemurs did not have anemia. This supports the conclusion that Otolemur species have moderate but not severe copper deficiency. Despite low diamine oxidase ceruloplasmin activity in Otolemurs, actual ceruloplasmin protein levels measured by immunoassay as well as the alternative ferroxidase II ceruloplasmin activity (11) were normal to elevated, which is expected given high serum copper levels (Table 1). The high serum copper, high serum copper/zinc ratio, and reversed copper/zinc ratio in liver support a chronic stress inflammatory response in even healthy Otolemurs (34). With high serum copper and ceruloplasmin protein levels, the relative depression of diamine oxidase activity and retention of ferroxidase II activity suggest alteration or loss of some ceruloplasmin copper enzymatic sites (11). This would be consistent with impairment of copper charging or posttranslational changes of some of the seven copper sites on ceruloplasmin. We measured serum vitamin E levels to exclude vitamin E deficiency as a cause of

116

SCHMECHEL ET AL.

TABLE 1

Maximum life span (years) Liver Steatosis Iron Copper Zinc Carnitine (Number) [CuZnSOD] [CytOX] [MnSOD] Ascorbate (Number) Serum Copper Zinc [Ceruloplasmin] (Number) Kidney Proximal tubules Steatosis NOS staining Mitochondrial abn CNS Dystrophic axons Spheroids Amyloidosis

Tupaia belangeri

Galago senegalensis

Otolemur crassicaudatus

Otolemur garnetti

12

12

18

22

Y 350 6 216 6.43 6 3.9 20.1 6 8.4 1.54 6 0.18 (9)

Y 302 6 101 3.89 6 0.8 33.9 6 9.3 0.59 6 0.18 (11)

Y 540 6 72 5.00 6 0.6 52.8 6 6.1 0.28 6 0.08 (44)

Y 1157 6 428 3.72 6 0.6 44.0 6 12 0.40 6 0.14 (27)

Y 2786 6 1343 9.19 6 5.2 105 6 54 1.03 6 0.33 (5)

Y 1493 6 1163 6.77 6 2.4 148 6 66 0.93 6 0.29 (3)

83.2 6 18.7 0.696 6 0.028 1.7 6 1.5 317 6 62 (6)

28.1 6 11.4 0.263 6 0.01 4.5 6 1.3 202 6 15 (14)

— 0.233 6 0.03 — 183 6 39 (6)

51.1 0.481 6 0.07 3.9 118 6 43 (4)

— — — 335 6 156 (3)

45 6 23 55 6 13 10 6 11 (5)

— — 57 6 24 (11)

141 6 12 100 6 15 52 6 6 (14)

159 6 30 107 6 23 48 6 10 (10)

— — —

85 6 12 92 6 19 81 6 25 (9)

6 N 6

6 N N

Y Y Y

Y Y Y

N N N

— — —

Occasionally N N

6 6 N

Y Y Y

Y Y Y

N N N

Occasionally N Oldest

— 0.443 6 .06 — — —

ceruloplasmin alteration or excessive ROS and found levels in Otolemur species of ca. 10 mg/liter (n 5 8 animals, normal range 5–15 mg/liter). The significance of the marginal copper and carnitine status of Otolemurs is supported by the finding that there is cardiac enlargement and hypertrophy in most adult animals. In 78 adult Otolemurs, 41 animals had heart weight/body weight ratios greater than 5.00 3 1024 (normally 4 3 1024 to 5 3 1024). In a series of 8 male Otolemurs aged 1–11 years, the 4 animals aged greater than 6 years had enlarged hypertrophic hearts. Heart weight correlated inversely with heart copper content (linear regression, P , 0.03, r2 5 56%) with the two most hypertrophied hearts having heart copper levels of ca. 4 µg/g wet weight.

Microcebus murinus 10

Eulemur f. collaris 35

Mitochondrial Abnormalities in Heart and Renal Proximal Tubule Epithelial Cells The lower values for liver carnitine and copper in Otolemur species suggested that they were most at risk for mitochondrial compromise. Ultrastructural analysis of liver from Otolemur did not show much evidence of abnormality except for the presence of micro- and macrosteatosis (Figs. 1C, 2A, and 2B) and increasing lipofuscin and mitochondrial autophagocytosis in the oldest animals. Stains for glycogen and general histology were unremarkable although there was age-related connective tissue portal bridging in all prosimians with high liver iron (Mallory trichrome stain). Ultrastructural analysis of left ventricular heart

FIG. 2. Mitochondrial morphology in liver, heart, and proximal renal tubule epithelial cells. (A, B) Section of adult Otolemur (13 years old, A; 6 years old, B) liver shows variable microsteatosis (A) and hepatocytes with some scattered age-related lipofuscin (arrowhead, B) and normal appearing mitochondria. Skeletal muscle and heart appeared also relatively normal although Otolemur heart often had multiple lines (two to three deep) and clustering of mitochondria between muscle fibers, suggesting increased mitochondrial volume (5-year-old Otolemur, C). Proximal renal tubular epithelial cells identified by brush border show normal mitochondrial morphology in aged M. murinus (D), although lipofuscin is common. Electron-dense mitochondrial inclusions are very common and obvious in Otolemur species particularly O. crassicaudatus (6 years old, E). Some scattered abnormal profiles consistent with degenerating mitochondria (F) are encountered in heart muscle of Otolemur (same animal as C). Higher magnification of proximal renal tubular epithelial cells in young Otolemur (1 year old) show small inclusions (G) compared to a high incidence of membranous whorls or inclusions (arrow) in older Otolemur (11 years old, H). Bar represents 0.5 µm (F). Original magnifications: (A, C) 77003; (B) 52503; (D, E) 12,4253; (F, G, and H) 36,7503.

CHOLINERGIC AXONAL DYSTROPHY IN PROSIMIANS

117

118

SCHMECHEL ET AL.

muscle showed evidence for mitochondrial hypertrophy with multiple lines of mitochondria or even clumps between many muscle fibers (Fig. 2C), and occasional profiles of abnormally dense mitochondria with condensed cristae (Fig. 2F) presumed to be degenerating mitochondria. No apparent excess fat was visualized in cardiac or skeletal muscle, and there were no ragged red fibers. Analysis of kidney tissue by routine histological stains often showed mild chronic tubulo-interstitial nephritis changes in the oldest animals and cystic changes in renal cortex, but typically no evidence for significant glomerular disease. Ultrastructural analysis showed relatively normal glomerular structure and basement membrane. In contrast, renal proximal tubule epithelial cells identified by brush border commonly showed widespread mitochondrial inclusion abnormalities in the two Otolemur species (Fig. 2E) and no significant abnormalities in other species (Fig. 2D). These abnormalities consisted of membranous whorls and electron-dense inclusions in mitochondria (Fig. 2H), which were minimally present in young Otolemur (Fig. 2G) and increased steadily with age (Fig. 2H). Mitochondrial inclusions were significantly more marked in O. crassicaudatus compared to O. garnetti. Mitochondrial abnormalities were not observed in glomerular cells, endothelial cells, or distal tubular epithelial cells. NADPH diaphorase staining of kidney tissue supported a marked species difference with increased diaphorase activity in the renal proximal tubule epithelial cells of Otolemur with mitochondrial abnormalities (Figs. 1E–1G) as well as marked fatty change visualized by oil red O staining (data not shown). Mitochondrial Abnormalities in Choroid Plexus Epithelial Cells and Pancreatic Islet Cells These two tissues were surveyed in Otolemur because of the similarity of choroid plexus epithelial cells to renal proximal tubular epithelial cells in terms of high mitochondrial volume density and metabolic load and because of the relationship of iron overload to pancreatic dysfunction (15). Both choroid plexus epithelial cells and pancreatic islet cells in Otolemur were strongly positive in NADPH-diaphorase staining, suggesting NOS induction of these tissues (8, 21). Choroid plexus epithelial cells commonly showed electrondense structures in young (Fig. 3A) and old Otolemur

(Fig. 3C). Closer inspection of these structures revealed that many represented profiles of degenerating mitochondria as identified by an outer double membrane or cristae-like membranes within (Figs. 3B and 3D). There were few intermediate forms between normal mitochondria and these clearly abnormal structures. Ependymal cells did not show this degree of change. In pancreatic islet cells, there were also frequent abnormal mitochondria with increased electron density of cristae, ballooning of matrix, and degenerating mitochondrial profiles (Figs. 3E, 3F, and 3G). The specificity of mitochondrial change in islet cells was supported by the lack of any significant abnormalities of mitochondria in pancreatic acinar cells unlike the acinar changes reported for severe dietary copper deficiency (12). Mitochondrial Abnormalities in Central Cholinergic Fibers and Cell Bodies (CH5-6) In the same Otolemurs with renal, pancreas, and choroid plexus mitochondrial abnormalities, significant changes in mitochondrial morphology and characteristics are observed in selected central cholinergic systems. For these studies, we utilized immunolocalization of manganese superoxide dismutase (MnSOD) as a convenient anatomical and functional marker for mitochondria (7, 39). For example, cholinergic cell bodies and centrally contained fibers of third cranial nerve show normal patterns of immunoreactivity for ChAT and the mitochondrial marker MnSOD in Otolemurs (Figs. 1I, 1J, and 4A), whereas nearby cholinergic cell bodies of PPN and laterodorsal tegmental nuclei (LDT) have extremely dark staining for MnSOD (Figs. 1K and 4B). In regions of cholinergic axonal terminals innervated by PPN such as dorsolateral geniculate body (dLGN), nucleus reticularis thalami (nRT), or basal forebrain, abnormally swollen cholinergic axons can be observed (Fig. 1L) again with abnormally large, very intensely immunoreactive MnSOD profiles present. Dystrophic cholinergic axons are easily visualized in nRT with ChAT staining (Fig. 4C) and less well with MnSOD localization (Fig. 4D). Cell bodies and dendrites of nRT neurons do not show this change in intensity or character of MnSOD staining. In nRT, dystrophic change is very selective for cholinergic fibers and specific immunolocalization shows no apparent abnormality of GABAergic or serotoninergic axons (data not shown). Most of these cholinergic dystrophic fibers

FIG. 3. Mitochondrial morphology in choroid plexus epithelial cells and pancreas islet cells. Choroid plexus epithelial cells identified by brush border had prominent electron-dense profiles of abnormal and degenerating mitochondria illustrated in young (5 years old, A) and older Otolemur (11 years old, C). These commonly encountered profiles could be confirmed as degenerating mitochondria by the presence of double membrane and cristae-like membrane fragments within the electron-dense interior (B, D). In pancreatic islet cells, there was a similar prominence of abnormal mitochondrial profiles in both young and old animals, shown in (E) and (F). In severely affected islet cells, there were marked mitochondrial changes with vacuolation, focal densities near intersections of cristae, and apparent enlargement (E). Other islet cells showed less marked changes and presented profiles more reminiscent of kidney and choroid plexus (G). Original magnifications: (A, C, E, F) 77003. (B, D, G) 36,7503.

CHOLINERGIC AXONAL DYSTROPHY IN PROSIMIANS

119

120

SCHMECHEL ET AL.

CHOLINERGIC AXONAL DYSTROPHY IN PROSIMIANS

are NADPH-diaphorase positive, supporting their identification as axon terminals of the LDT–PPN cholinergic complex of midbrain (42) (data not shown). Dystrophic fibers are rare or absent in other central cholinergic projections as habenular-interpenduncular pathway (CH7) and cranial nerve nuclei and tracts, as well as in the dense local circuit derived cholinergic innervation of striatum. When ChAT immunostained sections are inspected, the most obviously affected regions in young animals are nRT and dorsal thalamic nuclei such as dLGN (compare Figs. 4C, 4G, and 4I). This permits numerical characterization of density of abnormal dystrophic and normal cholinergic fibers in nRT as a way to compare different animals and species (Figs. 5A and 5B). Immunoelectronmicroscopy supports the identification of large dystrophic fibers in nRT as cholinergic and illustrates their abnormal morphology with a dense membranous interior and larger mitochondria than surrounding fibers and cells (Figs. 5C and 5D). A pilot survey of one animal revealed mitochondria in dystrophic cholinergic terminals to be 8100 6 3900 Å2 (n 5 22) compared to smaller mitochondria in surrounding axon terminals which averaged 4330 6 690 Å2 (n 5 48). An ultrastructral survey of large neurons in a region of PPN-LDT shows that many have numerous lipofuscin granules and other profiles consistent with degenerating mitochondria (Figs. 5E and 5F). These profiles may correlate with the increased MnSOD immunoreactivity of these neurons (Figs. 1I, 1K, 4A, and 4B). Mitochondrial Abnormalities Are Not Confined to the PPN-LDT Cholinergic System Using MnSOD immunoreactivity as a probe for abnormal mitochondria and swollen axons, it was common to observe distinct regions with dystrophic fibers or spheroids in other parts of the neuraxis including nucleus gracilis and cuneatus, deep cerebellar nuclei, central gray (Fig. 4C), molecular layer of dentate gyrus, and stratum lacunosum-moleculare of hippocampus (perforant pathway distribution), layer I of many neocortical regions (Fig. 4H), both ventral and dorsal striatum, and pallidum and anterior lateral hypothalamus (Fig. 4F). These changes were observed in the presence of widespread gliosis of both astrocytes and microglia with

121

MHC class II activation (these glia are strongly MnSOD immunoreactive) (34). In some regions, dystrophic cholinergic fibers were also observed. For example, dystrophic cholinergic fibers could be observed in layer I of neocortex or in hippocampus where MnSOD immunoreactive spheroids or fibers were observed (Figs. 4G, 4H, and 4I). These presumably derived from cholinergic neurons of basal forebrain (CH1-4). In many other regions such as terminal zones of the perforant pathway in hippocampus, striatal structures where dystrophic cholinergic fibers were rare, lateral hypothalamus, and dorsal column nuclei, MnSOD immunoreactive fibers and spheroids presumably represented dystrophic changes in other noncholinergic systems. Indeed, dystrophic TH- and 5HT-immunoreactive fibers were visualized in other forebrain regions (data not shown). Degree of Cholinergic Injury Is Species Specific and Closely Related to Systemic Factors The most marked cholinergic axonal dystrophy is observed in the two Otolemur species which are differentiated from the three other prosimian species and Tupaia by the presence of peripheral indices of stress inflammatory response (e.g., high serum copper, NOS induction in pancreatic islet cells, and proximal renal tubule epithelial cells) and possible carnitine and copper deficiency (Table 1). It is important to emphasize that Otolemurs did not appear sick, were reproductively active, and were living to lifespans exceeding those of feral animals. The availability of numerical estimates for cholinergic injury in a large number of animals as well as the indices of trace mineral and carnitine metabolism allowed testing the proposition that central cholinergic injury is closely associated with factors that might compromise mitochondrial function in cholinergic neurons and other tissues. In Fig. 6, we present graphs of the relationship of cholinergic axonal dystrophy in nRT to age and liver copper levels for O. garnetti. Density of dystrophic cholinergic axons in adult O. garnetti are not highly correlated with age (Fig. 6A). Increased cholinergic axonal dystrophy in nRT of O. garnetti is highly correlated with decreased liver copper levels (Fig. 6B; n 5 24 animals, P , 0.0001; r2 5 52% for linear regression). For O. crassicaudatus, liver copper declines steadily

FIG. 4. Dystrophic changes and spheroid formation in Otolemur. Comparison of MnSOD localization in noncholinergic brainstem neuron located in upper pons (A) and the more intense and larger punctate staining of presumptive cholinergic neuron located in PPN complex (B) of same Otolemur. In older Otolemurs, MnSOD immunoreactive spheroids were numerous in specific regions such as midbrain central gray (C). Marked dystrophic changes were typically observed in ChAT-immunoreactive cholinergic axon terminals innervating the nRT of older Otolemurs (7 years old, D). Semiadjacent section shows relatively fewer numbers of dystrophic fibers revealed by MnSOD staining without the presence of spheroids (E). In some regions such as anterolateral hypothalamus, both dystrophic fibers (arrowhead) and spheroids were detected by MnSOD immunolocalization (F). Dystrophic cholinergic axons were also observed in regions such as layer I of cerebral cortex (G) and various subfields of hippocampus, illustrated for CA4 (I). Again, MnSOD localization usually revealed fewer dystrophic fibers in these same regions (H), illustrated for an adjacent section of layer I (v indicates vessel). Original magnifications: (A, B) 6803; (C, D, E, G, H) 2503; (F) 4003.

122

SCHMECHEL ET AL.

CHOLINERGIC AXONAL DYSTROPHY IN PROSIMIANS

123

FIG. 6. (A) Cholinergic dystrophy in nRT vs Age. (B) Cholinergic dystrophy vs liver copper. (C) Cholinergic denervation in nRT. (D) Cholinergic denervation in dLGN.

with age and the cholinergic dystrophy in nRT is more closely correlated with age and liver carnitine level. Essentially no dystrophic fibers were visualized in nRT of M. murinus and T. belangeri with intermediate amounts in G. senegalensis. Cholinergic axonal dystrophy is accompanied by a progressive loss of the normal fine innervation of the nRT by cholinergic axons and terminals. Thus, on average 50% of original cholinergic innervation is lost at 14–15 years of age according to the regression line for O. garnetti (Fig. 6C; n 5 10 animals aged 4–22 years old, linear regression P , 0.0001, r2 5 85%) compared to 50% loss at 13–14 years of age for O. crassicaudatus (n 5 23 animals aged 0.5–15 years old; linear regression P , 0.005, r2 5 32%). The point of 50% denervation for both Otolemur species lies at roughly 70% of the maximum life span observed in captivity. The specificity of this observation in nRT is buttressed by the observation of no significant cholinergic denervation in striatum (O. crassicaudatus, n 5 7 animals) where few dystrophic cholinergic fibers are observed and no cholinergic denervation in nRT of T. belangeri where essentially no dystrophic cholinergic fibers are observed. In

G. senegalensis, there is 20–30% cholinergic denervation of the nRT in animals 8–9 years old, yielding a similar rate to Otolemur based on actual years of life, but a much slower rate based on expected maximal life span (Table 1). The identification of many of the affected cholinergic fibers in Otolemur species as axons of the midbrain cholinergic nuclei is supported by the finding that there is also a corresponding cholinergic denervation of the dorsolateral geniculate nucleus (dLGN) which reaches 50% denervation at age 14 years (Fig. 6D, subset of 10 animals aged 0.5–12 years; linear regression P , 0.00001, r2 5 96%). In a given animal, there is a strong correlation between degree of nRT and dLGN cholinergic denervation (n 5 4 animals; linear regression, P , 0.0001, r2 5 99%). DISCUSSION

The ultrastructural and immunocytochemical data in this report provide evidence that the marked cholinergic axonal dystrophy and denervation observed in captive Otolemurs involves early mitochondrial abnor-

FIG. 5. Illustration of the character and morphology of dystrophic cholinergic fibers in nRT from camera lucida drawings of normal fibers in young (A) and abnormal cholinergic fibers in old Otolemur (B). Bar represents 50 µm. ChAT immunoelectronmicroscopy of large dystrophic cholinergic fibers in nRT showing strong immunoreactivity of these large processes with possible cholesterol cleft crystal (arrowhead, C) and mitochondria that are generally larger and more dysmorphic (arrowhead, D) than those of surrounding normal tissue. Bar represents 2 µm. Original magnification, 77003. Many large neurons of the PPN complex showed extensive lipofuscin, but on closer examination many of these profiles had characteristics of degenerating mitochondria with internal linear membranous arrays reminiscent of cristae and focal electron-dense deposits. In most cases, only a single outer membrane was observed (arrowhead, E), but close inspection of some suggested the possible aggregation of a second putative inner membrane to electron-dense centers. Groupings often suggested clumped mitochondria (arrowhead, F). Bar represents 0.2 µm.

124

SCHMECHEL ET AL.

mality of the affected axons and the presence of degenerating mitochondrial profiles in parent cell bodies. The mitochondrial pathology observed in central cholinergic neurons is paralleled by mitochondrial pathology in selected cell classes of peripheral tissues and is specifically correlated with copper and carnitine deficiency demonstrated by biochemical methods. Cholinergic axonal dystrophy is easily demonstrated in prosimians using specific immunocytochemistry for ChAT to inspect the distal processes of the midbrain PPN-LDT projection to nRT and dorsal thalamus (34). We have observed dystrophic cholinergic axons in every captive wild-caught and colony bred adult animal, from two Otolemur species, that were autopsied. Cholinergic axonal dystrophy is a clinically ‘‘silent’’ process, beginning at 10% maximum life span. Colony-raised Otolemurs are reproductively fit to 12 years of age, are generally in good health until advanced age, and live to 20 years or more of age compared to maximal life spans of 7–10 years in the wild. At this time, we do not know whether wild Otolemurs would show cholinergic axonal dystrophy of this degree and do not know whether this process in captive Otolemurs may have detectable behavioral correlates as the animals age. Cholinergic axonal dystrophy in Otolemurs results in progressive denervation of targets of the midbrain PPN–LDT complex (CH5-6). Using the densely innervated thalamic target of nRT as index makes possible numerical estimates of the amount of cholinergic axonal dystrophy and associated denervation in a particular animal or species. Estimating for a time point of 50% of maximum life span, Otolemurs show nearly 50% loss of cholinergic innervation of nRT compared to ,20% loss for G. senegalensis Moholi and probably E. fulvus collaris, and almost no loss for T. belangeri and probably M. murinus. Thus, the degree of age-related cholinergic axonal dystrophy and associated central cholinergic denervation in prosimians and related species varies markedly. Cholinergic axonal dystrophy is not limited to prosimians and occurs during normal aging in many species including rodents, primates, and humans, in association with experimental lesions in animals, and in neurodegenerative diseases such as Alzheimer’s disease (AD), progressive supranuclear palsy (PSP), and Parkinson’s disease (PD) (6, 14, 50, 51). In humans, cholinergic axonal dystrophy can affect the cholinergic projections of the PPN–LDT or CH5-6 complex (14, 25, 50, 51) as well as the septal/diagonal band and nucleus basalis of Meynert complex (CH1-4) with associated cytoskeletal pathology in the neuronal cell bodies and processes (25, 43, 50, 51). Axonal dystrophy in AD also affects other projections of the diffuse ascending reticular activating system including the noradrenergic projections of the locus coeruleus (29). In prosimians, cholinergic axonal dystrophy begins in PPN–LDT sys-

tem, but does eventually involve the forebrain cholinergic projections at later ages. There is relative sparing even in the oldest animals of cholinergic neurons of the medial habenula, parabigeminal nucleus, striatum, and cranial nerves. Our previous report suggests a close relationship between the degree of cholinergic axonal dystrophy in prosimian species and the occurrence of reactive gliosis including astrocytosis and microgliosis with MHC class II activation (34). Extensive b-amyloidosis is observed in these species at age points corresponding to greater than 50% denervation of the nRT (34). The extensive degree of cholinergic dystrophy in Otolemurs represents pathological aging and fits into the larger neuropathological category of neuroaxonal dystrophy or spheroid formation. Neuroaxonal dystrophy with morphological changes of focal axonal swelling and spheroid formation is rather nonspecific for nervous system injury (20, 37, 47). Axonal dystrophy is described in normal aging, in human diseases such as infantile neuroaxonal dystrophy and vitamin E deficiency, in association with cancer, and in a variety of late-onset human neurodegenerative diseases such as PD, PSP, and AD (6, 14, 29, 47). In this report, we can more specifically associate cholinergic axonal dystrophy even at its earliest time points with mitochondrial pathology. Our observations in Otolemurs of the presence in PPN–LDT neurons of lipofuscin and profiles of degenerating mitochondria, the demonstration that spheroids are MnSOD immunoreactive, and the demonstration of abnormal dystrophic fibers by MnSOD immunoreactivity support an early abnormality of mitochondria even in axons with minimal swelling or morphological abnormalities. While the PPN–LDT cholinergic neurons are a ‘‘sentinel’’ system in prosimians, axonal dystrophy also affects noncholinergic projections. The use of MnSOD immunoreactivity as a marker for both normal and abnormal mitochondria permitted the detection of abnormal noncholinergic cells, axons, and spheroids (presumably degraded or even phagocytosed axonal swellings and neurites) in a number of other defined brain areas besides nRT. In particular, there was evidence for damage to perforant pathway projections, layer I projections, and subcortical projections to hypothalamus, central gray, and dorsal and ventral striatum and pallidum. Thus, there is evidence for widespread selective axonal dystrophy and mitochondrial pathology in Otolemurs in addition to the sentinel cholinergic projections of PPN-LDT. Mitochondrial pathology in nervous system aging is well described. Early studies of nervous system aging described a mitochondrial etiology for some lipofuscin production and spheroids or corpora amylacea (16, 35). Some mitochondrial injury occurs during normal aging of primate brain (5). Mitochondrial mtDNA alterations

CHOLINERGIC AXONAL DYSTROPHY IN PROSIMIANS

may occur during AD, possibly as a result of oxidative damage to cells (9, 23, 38). Recently, a mtDNA variant was described as a risk or modulating factor in lateonset AD (19). Inherited or acquired mtDNA defects may be an important factor in human neurodegenerative diseases (44). Likewise, somatic mutations and their related proteins may affect directly or indirectly mitochondrial function and integrity (2, 46). Mitochondrial pathology in neurons in Otolemurs is accompanied by anatomical evidence for selective mitochondrial injury in other nonneuronal cell classes such as renal proximal tubule epithelial cells, pancreatic islet cells, and choroid epithelial cells in these same animals. Interestingly, cholinergic neurons of the CH5-6 group normally express nitric oxide synthetase (NOS) (10, 42) and all of the other three NOS-inducible cell classes (8, 21) above in affected animals were NOSpositive by diaphorase staining. All have high metabolic activity and high mitochondrial volume density compared to liver, which was relatively unaffected. This pattern of tissue involvement is different from some mitochondrial diseases, but similar to the pattern observed in the expression of mutant Cu,Zn-SOD in the transgenic animal model of ALS (46). The marked degree of axonal dystrophy and mitochondrial pathology in Otolemurs may relate directly to disordered copper and carnitine metabolism. The presence of mitochondrial abnormalities in peripheral tissues is closely associated with low values for liver copper, cuproenzymes, and carnitine in the same rank ordering as the extent of central cholinergic axonal dystrophy (i.e., Otolemur . Galago .. Microcebus, Tupaia, E. fulvus collaris). The close relationship of marginal copper status to this central process is supported by the correlation of increased cholinergic axonal dystrophy with low liver copper levels demonstrated for O. garnetti (Fig. 6B). There is biochemical evidence for relative copper deficiency in Otolemurs compared to other prosimian species based on low liver cytochrome oxidase and Cu,Zn-SOD activities and depressed diamine oxidase activity of ceruloplasmin (Table 1). In addition, there is the presence of abnormal mitochondria and cardiac myopathy. Copper deficiency can cause or possibly enhance mitochondrial dysfunction and pathology (30, 31). In the context of prosimian iron overload (1, 15), it is interesting to note that abnormalities or deficits of ceruloplasmin can lead to hemosiderosis (18, 48, 49). In the case of Otolemur, copper deficiency must be moderate and not severe since there is no anemia and the animals still generate high serum copper levels. Moreover, the high serum copper and apparent NOS induction in pancreas, kidney, and choroid plexus also support the concept that captive Otolemurs have chronic activation of the stress inflammatory response system (8, 31, 45) despite apparent good health and an absence of overt inflammatory

125

or infectious disease. Abnormal ceruloplasmin in this setting may not adequately deliver copper to peripheral target tissues. Moreover, abnormal ceruloplasmin and/or high serum copper may contribute to ROS formation and tissue injury. The other factor to be considered in the Otolemur syndrome of cholinergic axonal dystrophy and mitochondrial pathology is low carnitine stores. We have not identified a specific disorder of carnitine metabolism at this point, and screens for abnormal acylcarnitines, ascorbic acid deficiency, and organic aciduria are negative. Significant carnitine deficiency could impact fatty acid metabolism and predispose to cardiomyopathy, hepatic and brain dysfunction, and mitochondrial abnormalities (24, 41). Low carnitine stores in Otolemur could also represent a secondary effect of mild to moderate renal proximal tubule pathology with a partial Fanconi’s syndrome (4, 17, 24). Our present hypothesis is that carnitine abnormalities are secondary to mitochondrial pathology and associated renal pathology with presumed excessive renal carnitine loss. Studies of carnitine clearance, phosphate excretion, and amino acid excretion are underway to test this hypothesis. The pertinence of the prosimian model to human CNS aging is supported by the observation that prosimian species have a human pattern of neuronal apoE content and that the Otolemur species with the greatest degree of mitochondrial and cholinergic abnormalities are precisely those with an onset of b-amyloidosis at 50% of maximum life span (near the time point for 50% loss of cholinergic innervation of nRT) (34). We suggest that the prosimian model could mean that abnormal copper status might be associated with this same pattern of mitochondrial injury in subhuman primates and man. Gliosis with sequestration of copper stores, production of abnormal cuproproteins with loss of available copper for mitochondrial proteins, and other similar processes could result in those cells with greatest mitochondrial content being essentially copper deficient in the midst of other cells not so compromised. Mitochondrial injury and degeneration might then further compromise the situation by loss of copper stores, further tissue injury, and disruption of mtDNA by reactive oxygen species and so further drive this process. We have not excluded an inherited genetic factor in Otolemurs that might produce this syndrome such as an mtDNA defect or a defect in carnitine or copper metabolism. If there is an inherited genetic factor, this would be a species-wide trait since we observed the syndrome in two subspecies of Otolemur with both wild-caught and colony bred individuals, as well as a milder syndrome in G. senegalensis. Whether this syndrome is present in feral Otolemurs is unknown. Moreover, captivity-related environmentally induced

126

SCHMECHEL ET AL.

alterations in copper or carnitine status could conceivably generate new defects in maternal mtDNA that would then be transmissible to subsequent generations. We currently have some evidence for enhanced cholinergic dystrophy in some maternal lineages in O. crassicaudatus, but note that all lineages and both colony bred and wild caught animals show significant damage. Current research is directed at better delineating the abnormalities of copper and carnitine metabolism in these species with a special emphasis on environmental or dietary manipulations that might lessen the severity of cholinergic injury and mitochondrial pathology.

7.

8.

9.

10.

ACKNOWLEDGMENTS 11. D.E.S. expresses deep gratitude to Dr. Milton Brightman for his encouragement, teaching, and joyful approach to science that D.E.S. was privileged to receive during his stay at the National Institutes of Health (PRAT program, NIGMS/USPHS). We also thank Susan Reeves, Stephen Conlon, and Henry Estrada for excellent technical assistance. Much of the material was generated from anatomical experiments carried out in the laboratories of Dr. Irving T. Diamond and Dr. David Fitzpatrick with the participation of Dr. Michael Conley and Dr. Richard Penny. John Tsavaras helped immensely with the initial phases of measuring trace minerals in these animals. The assistance of many staff veterinarians at Duke Vivarium and DUPC is gratefully recognized. The gift of MnSOD antibody from Dr. Chang is gratefully acknowledged. This long-term study was supported by a Career Development Award from the VA, R-01 grants under NIA Alzheimer’s Disease Center Grant AG-05128, and NIEHS IAA Y01-ES-40290. The assistance of Compuchem, RTP, NC, and numerous private gifts to the Joseph and Kathleen Bryan ADRC and Duke University are also gratefully acknowledged. Finally, a great personal debt is acknowledged to D.L.S., D.A.S., F.W.S., and E.R.W.

12.

13.

14.

15. 16.

REFERENCES 17. 1.

2.

3.

4.

5.

6.

ABRAMSON, A. 1996. ‘Silent Disease’ in Captive Prosimians: Carnitine Deficiency and Iron Overload. Senior Honors Thesis, Duke University. ASKANAS, V., J. MCFERRIN, S. BAQUE, R. B. ALVAREZ, E. SARKOZI, AND W. K. ENGEL. 1996. Transfer of beta-amyloid precursor protein using adenovirus vector causes mitochondrial abnormalities in cultured normal human muscle. Proc. Natl. Acad. Sci. USA 93: 1314–1319. BARZANTI, V., M. BATTINO, A. BARACCA, M. CAVAZZONI, M. COCCHI, R. NOBLE, M. MARANESI, E. TURCHETTO, AND G. LENAZ. 1994. The effect of dietary lipid changes on the fatty acid composition and function of liver, heart, and brain mitochondria in the rat at different ages. Br. J. Nutrit. 71: 193–202. BERNARDINI, I., W. B., RIZZO, M. DALAKAS, J. BERNAR, AND W. A. GAHL. 1985. Plasma and muscle free carnitine deficiency due to renal Fanconi syndrome. J. Clin. Invest. 75: 1124–1130. BOWLING, A. C., E. M. MUTISYA, L. C. WALKER, D. L. PRICE, L. C. CORK, AND M. F. BEAL. 1993. Age-dependent impairment of mitochondrial function in primate brain. J. Neurochem. 60: 1964–1967. BRANDEL, J. P., E. C. HIRSCH, S. MALESSA, C. DUYCKAERTS, P. CERVERA, AND Y. AGID. 1991. Differential vulnerability of cholinergic projections to the mediodorsal nucleus of the thalamus in

18.

19.

20.

21.

22.

23.

24.

senile dementia of Alzheimer type and progressive supranuclear palsy. Neuroscience 41: 25–31. CHANG, L. Y., B. H. KANG, J. W. SLOT, R. VINCENT, AND J. D. CRAPO. 1995. Immunocytochemical localization of the site of superoxide dismutase induction by hyperoxia in rat lung. Lab. Invest. 73: 29–39. CORBETT, J. A., D. L., WONG, T. P. MISKO, W. ZHAO, B. F. HICKEY, AND M. L. MCDANIEL. 1993. Nitric oxide mediates IL-1-betainduced islet dysfunction and destruction: Protection by dexamethasone. Autoimmunity 15: 145–153. CORRAL-DEBRINSKI, M., T. HORTON, M. T. LOTT, J. M. SCOFFNER, A. C. MCKEE, M. F. BEAL, B. H. GRAHAM, AND D. C. WALLACE. 1994. Marked changes in mitochondrial DNA deletion levels in Alzheimer brains. Genomics 23: 471–476. DAWSON, T. M., D. S. BREDT, M. FOTUHI, P. M. HWANG, AND S. H. SYNDER. 1991. Nitric oxide synthase and neuronal NADPH diaphorase are identical in brain and peripheral tissues. Proc. Natl. Acad. Sci. USA 88: 7797–7801. EVANS, P. J., A. BOMFORD, AND B. HALLIWELL. 1989. Nonceruloplasmin copper and ferroxidase activity in mammalian serum: Ferroxidase activity and phenanthroline-detectable copper in human serum in Wilson’s disease. Free Radical Res. Commun. 7: 55–62. FELL, B. F., T. P. KING, AND N. T. DAVIES. 1982. Pancreatic atrophy in copper deficient rats: Histochemical and ultrastructural evidence of a selective effect on acinar cells. Histochem. J. 14: 665–680. FITZPATRICK, D., G. R. PENNY, AND D. E. SCHMECHEL. 1984. Glutamic acid decarboxylase-immunoreactive neurons and terminals in the lateral geniculate nucleus of the cat. J. Neurosci. 4: 1809–1829. GAYKEMA, R. P., C. NYAKAS, E. HORVATH, L. B. HERSH, C. MAJTENYI, AND P. G. LUITEN. 1992. Cholinergic fiber aberrations in nucleus basalis lesioned rat and Alzheimer’s disease. Neurobiol. Aging 13: 441–448. GONZALES, J., K. BENIRSCHKE, P. SALTMAN, J. ROBERETS, AND P. T. ROBINSON. 1984. Hemosiderosis in lemurs. Zoo. Biol. 3: 255–265. GOPINATH, G., AND P. GLEES. 1974. Mitochondrial genesis of lipofuscin in the mesencephalic nucleus of the vth nerve of aged rats. Acta Anat. 89: 14–20. GUDER, W. G., AND S. WAGNER. 1990. The role of the kidney in carnitine metabolism. J. Clin. Chem. Clin. Biochem. 78: 347– 350. HARRIS, Z. L., Y. TAKAHASI, H. MIYAJIMA, M. SERIZAWA, R. T. MACGILLIVRAY, AND J. D. GITLIN. 1995. Aceruloplasminemia: Molecular characterization of this disorder of iron metabolism. Proc. Natl. Acad. Sci. USA 92: 2539–2543. HUTCHIN, T., AND G. CORTOPASSI. 1995. A mitochondrial clone is associated with increased risk for Alzheimer disease. Proc. Natl. Acad. Sci. USA 92: 6892–6895. JELLINGER, K., AND A. JIRASEK. 1971. Neuroaxonal dystrophy in man: Character and natural history. Acta Neuropathol. 5(Suppl 5): 3–16. MARKEWITZ, B. A., J. R. MICHAEL, AND D. E. ROHEN. Cytokine induced expression of a nitric oxide synthase in rat renal tubule cells. J. Clin. Invest. 91: 2138–2143. MCCORD, J. M., AND I. J. FRIDOVICH. 1969. Superoxide dismutase: An enzymatic function for erythrocuprein (hemocuprein). J. Biol. Chem. 244: 6049. MECOCCI, P., U. MACGARVEY, AND M. F. BEAL. 1994. Oxidative damage to mitochondrial DNA is increased in Alzheimer’s disease. Ann. Neurol. 36: 747–751. MIYAGAWA, J., M. KUWAJIMA, T. HANAFUSA, K. OZAKI, H. FUJIMURA, A. ONO, R. UENAKA, I. NARAMA, T. OUE, K. YAMAMOTO, et

CHOLINERGIC AXONAL DYSTROPHY IN PROSIMIANS

25.

26.

27.

28.

29.

30.

31.

32. 33.

34.

35.

36.

37.

38.

al. 1995. Mitochondrial abnormalities of muscle tissue in mice with juvenile visceral steatosis associated with systemic carnitine deficiency. Virch. Archiv. 426: 271–279. MUFSON, E. J., D. C. MASH, AND L. B. HERSH. 1988. Neurofibrillary tangles in cholinergic pedunculopontine neurons in Alzheimer’s disease. Ann. Neurol. 24: 623–629. NGUYEN-LEGROS, J., J. BIZOT, M. BOLESSE, AND J. P. PULICANI. 1980. ‘‘Noir de diaminobenzidine’’: Une nouvelle methode histochimique de revelation du fer exogene. Histochem. 66: 239–244. OERTEL, W. H., A. M. GRAYBIEL, E. MUGNAINI, R. P. ELDE, D. E. SCHMECHEL, AND I. J. KOPIN. 1983. Co-existence of glutamic acid decarboxylase immunoreactivity and somatostatin-like immunoreactivity in neurons of the feline nucleus reticularis thalami. J. Neurosci. 3: 1322–1332. PARVIN, R., AND S. V. PANDE. 1977. Microdetermination of (2) carnitine and carnitine acetyl transferase. Anal. Biochem. 79: 190–201. POWERS, R. E., R. G. STRUBLE, M. F. CASANOVA, D. T. O’CONNOR, C. A. KITT, AND D. L. PRICE. 1988. Innervation of human hippocampus by noradrenergic systems: Normal anatomy and structural abnormalities in aging and Alzheimer’s disease. Neuroscience 25: 401–417. PROHASKA, J. R. 1991. Changes in Cu,Zn-superoxide dismutase, cytochrome c oxidase, glutathione peroxidase and glutathione transferase activities in copper deficient mice and rats. J. Nutr. 121: 355–363. PROHASKA, J. R., AND W. W. WELLS. 1975. Copper deficiency in the developing rat brain: evidence for abnormal mitochondria. J. Neurochem. 25: 221–228. REBOUCHE, C. J. 1991. Ascorbic acid and carnitine biosynthesis. Am. J. Clin. Nutr. 54 (6 Suppl): 1147S–1152S. REBOUCHE, C. J. 1995. The ability of guinea pigs to synthesize carnitine at a normal rate from epsilon-N-trimethyllysine or gamma-butyrobetaine in vivo is not compromised by experimental vitamin C deficiency. Metabolism 44: 624–629. SCHMECHEL, D. E., M. O. TILLER, P. TONG, M. MCSWAIN, S.-H. HAN, R. ANGE, D. S. BURKHART, AND M. K. IZARD. 1996. Pattern of apolipoprotein E immunoreactivity during brain aging. In Apolipoprotein E and Alzheimer’s Disease (A. D. Roses, K. H. Weisgraber, and Y. Christen, Eds.), pp. 29–48. Springer-Verlag, Berlin. SCHIPPER, H. M., AND S. CISSE. 1995. Mitochondrial constituents of corpora amylacea and autofluorescent astrocytic inclusions in senescent human brain. Glia 14: 55–64. SCHOSINSKY, K. H., H. P. LEHMANN, AND M. F. BEELER. 1974. Measurement of ceruloplasmin from its oxidase activity in serum by use of o-dianisidine dihydrochloride. Clin. Chem. 20: 1156–1563. SEITELBERGER, F. 1971. Neuropathological conditions related to neuroaxonal dystrophy. Acta Neuropathol. (Berlin) Suppl. 5: 17–29. SIMONIAN, N. A., AND B. T. HYMAN. 1994. Functional alterations in Alzheimer’s disease: Selective loss of mitochondrial-encoded

39.

40.

41.

42.

43.

44.

45.

46.

47.

48.

49.

50.

51.

127

cytochrome oxidase mRNA in the hippocampal formation. J. Neuropathol. Exp. Neurol. 53: 508–512. SLOT, J. W., H. J. GEUZE, B. A. FREEMAN, AND J. D. CRAPO. 1986. Intracellular localization of the copper-zinc and manganese superoxide dismutases in rat liver parenchymal cells. Lab. Invest. 55: 363–371. SMITH, L. 1974. Spectrophotometric assay of cytochrome c oxidase. In Methods of Biochemical Analysis (D. Glick, Ed.), Vol. 11, p. 427. Wiley, New York.. STANLEY, C. A. 1979. New genetic defects in mitochondrial fatty acid metabolism and carnitine deficiency. Adv. Pediatr. 34: 59–88. SUGAYA, K., AND M. MCKINNEY. 1994. Nitric oxide synthase gene expression in cholinergic neurons in the rat brain examined by combined immunocytochemistry and in situ hybridization histochemistry. Mol. Brain Res. 23: 111–125. TOURTELLOTTE, W. G., G. W. VAN HOESEN, B. T. HYMAN, R. K. TIKOO, AND A. R. DAMASIO. 1990. Alz-50 immunoreactivity in the thalamic reticular nucleus in Alzheimer’s disease. Brain Res. 515: 227–234. WALLACE, D. C., J. M. SHOFFNER, I. TROUNCE, M. D. BROWN, S. W. BALLINGER, M. CORRAL-DEBRINSKI, T. HORTON, A. S. JUN, AND M. T. LOTT. 1995. Mitochondrial DNA mutations in human degenerative diseases and aging. Biochem. Biophys. Acta 1271: 141–151. WARD, W. F., A. MOLTENI, C., TS’AO, AND M. ISCHIROPOULOS. 1989. Serum copper as an index of experimental lung injury. Adv. Exper. Med. Biol. 258: 787–302. WONG, P. C., C. A. PARDO, D. R. BORCHETT, M. K. LEE, N. G. COPELAND, N. A. JENKINS, S. S. SISODIA, D. W. CLEVELAND, AND D. L. PRICE. 1995. An adverse property of familial ALS-linked SOD1 mutation causes motor neuron disease characterized by vacuolar degeneration of mitochondria. Neuron 14: 1105–1114. YAGASHITA, S. 1978. Morphological investigations on axonal swellnigs and spheroids in various human disease. Virch. Archiv. A Pathol. Anat. Hist. 378: 181–197. YOSHIDA, K., K. FURIHATA, S. TAKEDA, A. NAKAMURA, K. YAMAMOTO, H. MORITA, S. HIYAMUTA, S. IKEDA, N. SHIMIZU, AND N. YANAGISAWA. 1995. A mutation in the ceruloplasmin gene is associated with systemic hemosiderosis in humans. Nature Genet. 9: 267–272. YUAN, D. S., R. STEARMAN, A. DANCIS, T. DUNN, T. BEELER, AND R. D. KLAUSNER. 1995. The Menkes/Wilson disease gene homologue in yeast provides copper to a ceruloplasmin-like oxidase required for iron uptake. Proc. Natl. Acad. Sci. USA 92: 2632–2636. ZWEIG, R. M., W. R. JANKEL, J. C. HEDREEN, R. MAYEUX, AND D. L. PRICE. 1989. The pedunculopontine nucleus in Parkinson’s disease. Ann. Neurol. 26: 41–46. ZWEIG, R. M., P. J. WHITEHOUSE, M. F. CASANOVA, L. C. WALKER, W. R. JANKEL, AND D. L. PRICE. 1987. Loss of pedunculopontine neurons in progressive supranuclear palsy. Ann. Neurol. 22: 18–25.