Mutagenesis Advance Access published online on August 20, 2007
Mutagenesis, doi:10.1093/mutage/gem029
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Published by Oxford University Press on behalf of the UK Environmental Mutagen Society.
Buccal micronucleus cytome biomarkers may be associated with Alzheimer's disease
1CSIRO Division of Human Nutrition, 13 Kintore Avenue, PO Box 10041 Adelaide BC, Adelaide, SA 5000, Australia 2Discipline of Physiology, School of Molecular and Biomedical Sciences, The University of Adelaide, Adelaide, SA 5005, Australia 3Department of Internal Medicine, Royal Adelaide Hospital, North Terrace, Adelaide, 5000 4Royal Adelaide Hospital, Adelaide, Australia
Alzheimer's disease (AD) is a progressive degenerative disorder of the brain and is the commonest form of dementia. A buccal cytome assay was used to measure ratios of buccal cell populations and micronuclei in clinically diagnosed Alzheimer's patients compared to age and gender-matched controls. Frequencies of basal cells (P < 0.0001), condensed chromatin cells (P < 0.0001) and karyorrhectic cells (P < 0.0001) were found to be significantly lower in Alzheimer's patients. These changes may reflect alterations in the cellular kinetics or structural profile of the buccal mucosa, and may be useful as potential biomarkers in identifying individuals with a high risk of developing AD. The odds ratio of being diagnosed with AD for those individuals with a basal cell plus karyorrhectic cell frequency <41 per 1000 cells is 140, with a specificity of 97% and sensitivity of 82%. These promising results need to be replicated in larger studies and in cohorts of other neurodegenerative disorders to determine specificity of changes to Alzheimer's patients.
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Introduction |
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Alzheimer's disease (AD) is a neurodegenerative disorder that is characterized clinically by cognitive decline, memory loss, visuospatial and language impairment and is the commonest form of dementia (1
–4). Regions of the brain that are involved in short-term memory and learning such as the temporal and frontal lobes are impaired as a result of neuronal loss and the breakdown of the neuronal synaptic connections (5). The disease is histopathologically confirmed at post-mortem within the brain by the presence of neurofibrillary tangles and amyloid plaques (6,7). The greatest risk factor for contracting the disease is advancing age and risk is significantly increased beyond the age of 70 years (1,8,9). To date, the global prevalence is estimated to be 24.3 million with that number expected to triple within the next 30 years. At least 4.9 million new cases are reported annually with a predicted prevalence worldwide of 80 million people suffering from the disease by 2040 (9,10).
Currently, clinical diagnosis of AD is based upon criteria of cognitive impairment and behavioural changes (11). One such diagnostic tool is the mini mental state examination (MMSE), which allows a quantitative measurement of cognition to be determined (12,13). However, the diagnosis for having AD is only between 60 and 70% accurate when using these criteria and can only be conclusively confirmed after histopathological investigation (11).
Biomarkers that may identify individuals who are at an early stage of AD would be useful as this would allow timely preventative intervention. There is an increasing interest in the evaluation of chromosome damage markers within the somatic cells of neurodegeneration patients (14–16). Genome damage could lead to altered gene dosage and gene expression as well as contribute to the risk of accelerated cell death in neuronal tissue (17). Genomic instability markers such as micronuclei (MNs), which are biomarkers of chromosome malsegregation and breaks, have been shown to be elevated within the peripheral blood lymphocytes and fibroblasts of Alzheimer's patients (14,16,18).
The aim of this study was to assess whether MNs and other parameters of genome damage and cell death in the buccal mucosa could be used as a non-invasive method of identifying those at risk of developing AD.
In effect we adopted a cytome approach in which all the various cell types and their ratios were quantified to identify which parameters, if any, were most strongly associated with AD. A flow diagram illustrating plausible cellular relationships of the buccal cytome assay is shown in Figure 1.
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Methods |
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Recruitment and characteristics of participants
Approval for this study was obtained from CSIRO Human Nutrition, Adelaide University and Ramsay Healthcare Ethics Committee's. Twenty-six older controls (age 66–75 years) and 54 clinically diagnosed Alzheimer's patients (age 58–93 years) not receiving anti-folate therapy, cancer treatment or having any family history of AD were recruited for the study. Alzheimer's patients were recruited at the College Grove Private Hospital, Walkerville, Adelaide, South Australia, following their initial diagnosis and prior to commencement of therapy. Diagnosis of AD was made by clinicians according to the criteria outlined by the National Institute of Neurological and Communicative Disorders and Stroke-Alzheimer's Disease and related Disorders Association (NINCDS–AD&DA) (13
), which are the well recognized standards used in all clinical trials. Volunteers did not receive any remuneration for their participation. Alzheimer's patients were separated into two distinct groups. One group was age matched to the control, whereas the second group was classified as an older Alzheimer's cohort. Age, gender and MMSE scores of the cohorts are shown in Table I. Gender ratio differences between the groups were not significant (chi square P > 0.05). MMSE scores between the two AD groups were not significantly different, P > 0.05. The age of the control group compared to the younger AD group was not significantly different but the latter group had a significantly lower age relative to the older AD group, P < 0.0001. MMSE scores were not available for the controls. The controls were ‘normal’ functioning healthy individuals, who self-volunteered and consented to the study and did not report a history of cognitive impairment.
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Cell sampling and preparation
Buccal cells (BCs) were collected from consented volunteers by the author P.T. Cells were collected using a modified version of the method used by Beliën et al. (19
). Prior to BC collection the mouth was rinsed thoroughly with water to remove any unwanted debris. Small-headed toothbrushes (Supply SA, code 85300012) were rotated 20 times in a circular motion against the inside of the cheek, starting from a central point and gradually increasing in circumference to produce an outward spiral effect. Both cheeks were sampled using separate brushes. The heads of the brushes were individually placed into separate 30-ml yellow top containers (Sarstedt code 60.9922.918) containing BC buffer [0.01 M Tris–HCl (Sigma T-3253), 0.1 M EDTA tetrasodium salt (Sigma E5391), 0.02 M sodium chloride (Sigma S5886)] at pH 7.0 and agitated to dislodge the cells. Cells from both right and left cheeks were transferred into separate TV-10 centrifuge tubes (Sarstedt, code 60.9921.829) and spun for 10 min at 1500 rpm (MSE Mistral 2000). Supernatant was removed and replaced with 10 ml of fresh BC buffer. The BC buffer helps to inactivate endogenous DNAase and aids in removing bacteria that may complicate scoring. Cells were spun and washed twice more, with a final volume of 5 ml of BC buffer being added to the cells. The cell suspension was vortexed and then homogenized for 2 min in a hand homogenizer (Wheaton Scientific 0.1- to 0.15-mm gauge) to increase the number of single cells in suspension. Left and right cell populations were pooled into a 30-ml yellow top container and drawn into a syringe with a 21-G gauge needle and expelled to encourage cellular separation. Cells were passed into a TV-10 tube through a 100-µm nylon filter (Millipore, code MILNYH02500) held in a swinex filter (Millipore, code MILSX0002500) to remove large aggregates of unseparated cells that hinder slide preparation and cell analysis. Cells were further spun at 1500 rpm for 10 min and the supernatant was removed. Cells were re-suspended in 1 ml of BC buffer and the cell concentration was determined by a Coulter counter (Beckman Coulter Model ZB1; settings: attenuation, 1; threshold, 8; aperture, 1/4; manometer, 0.5). Cell suspensions were prepared containing 80 000 cells/ml after initial readings from a 1:50 dilution (300 µl/15 ml isoton). Dimethyl sulphoxide (50 µl/ml; Sigma 2650) was added to help clarify cellular boundaries by further separating the cells. Cell suspension (120 µl) was added to cytospin cups and spun at 600 rpm for 5 min in a cytocentrifuge (Shandon cytospin 3). Slides containing two spots of cells were air-dried for 10 min and then fixed in ethanol:acetic acid (3:1) for 10 min. Slides were air-dried for 10 min prior to staining.
Feulgen staining
Fixed slides were treated for 1 min each in 50 and 20% ethanol and then washed for 2 min in deionized water generated from a Milli-q water purification system (Millipore Australia Pty Ltd, New South Wales, Australia). Slides were treated in 5 M hydrochloric acid for 30 min and then washed in running tap water for 3 min. Slides were drained but not allowed to dry out before being treated in room temperature Schiff's reagent (Sigma 3259016) in the dark for 60 min. Slides were washed in running tap water for 5 min and rinsed well in deionized water for 1 min. Slides were stained for 30 sec in 0.2% light green (Sigma L-1886) and rinsed well in deionized water for 2 min. Slides were allowed to air-dry, covered with No. 1 coverslips (Crown Scientific, South Australia) and mounted in DePeX (BDH 361254D). Nuclei and MNs are stained magenta, while the cytoplasm appears green. Slides were scored using a Nikon E600 microscope equipped with a triple-band filter (4,6-diamidino-2-phenylindole, fluorescein isothiocyanate and rhodamine) at x1000 magnification. Cells containing MNs on bright field can be confirmed as being positive by examining the cells under fluorescence. The incidence of false positives can be minimized as DNA material such as nuclei and MNs fluoresce when viewed under fluorescence with a far-red filter (Figure 2).
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Scoring criteria for buccal cytome assay
The various distinct populations used in the buccal cytome assay were determined based on criteria outlined by Tolbert et al. (20
). These criteria are intended to classify BCs into categories that distinguish between ‘normal’ cells and cells that are considered ‘abnormal', based on nuclear morphology. These abnormal nuclear morphologies are thought to be indicative of DNA damage or cell death. Photographic images showing distinct cell populations as scored in the buccal cytome assay are shown in Figure 3. Detailed descriptions of the various cell types are given below.
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Basal cells. These are the cells from the basal layer (Figure 3A). The nuclear to cytoplasm ratio is larger than that in differentiated BCs that are derived from basal cells. Basal cells have a uniformly stained nucleus and they are smaller in size when compared to differentiated BCs. Basal cells can contain MNs and were scored in the assay.
Normal differentiated cells. These cells have a uniformly stained nucleus that is usually oval or round in shape (Figure 3B). They are distinguished from basal cells by their larger size and by a smaller nuclear to cytoplasmic ratio. No other DNA-containing structures apart from the nucleus are observed in these cells. These cells are considered to be terminally differentiated relative to basal cells because no mitotic cells are observed in this population.
Cells with MNs.
These cells are characterized by the presence of both a main nucleus and one or more smaller nuclei called MNs (Figure 3A and C). The MNs are usually round or oval in shape and their diameter may range between 1/3 and 1/16 the diameter of the main nucleus. Cells with MNs usually contain only one micronucleus. It is possible but rare to find cells with more than six MNs. The nuclei in micronucleated cells may have the morphology of normal cells or that of dying cells (i.e. condensed chromatin cells). The MNs must be located within the cytoplasm of the cells. The presence of MNs is indicative of chromosome loss or fragmentation occurring during previous nuclear division (21). MNs were scored only in basal and differentiated cells with uniformly stained nuclei. Cells with condensed chromatin or karyorrhectic cells were not scored for MNs.
Cells with nuclear buds.
These cells have nuclei with an apparent sharp constriction at one end of the nucleus suggestive of a budding process, i.e. elimination of nuclear material by budding (Figure 3D). The nuclear bud and the nucleus are usually in very close proximity and are apparently attached to each other. The nuclear bud has the same morphology and staining properties as the nucleus; however, its diameter may range from a half to quarter of that of the main nucleus. The mechanism leading to this morphology is not known but it may be due to elimination of amplified DNA or DNA repair complexes (22–24).
Binucleated differentiated cells. These cells have two nuclei instead of one (Figure 3E). The nuclei are usually very close to each other and may be touching. The nuclei usually have the same morphology as that observed in normal cells. The significance of these cells is unknown but they may be indicative of failed cytokinesis following the last nuclear division.
Condensed chromatin cells. These cells have nuclei with regions of condensed or aggregated chromatin exhibiting a speckled or striated nuclear pattern (Figure 3F). In these cells, it is apparent that chromatin is aggregating in some regions of the nucleus while being lost in other areas. When chromatin aggregation is extensive, the nucleus may appear to be fragmenting. These cells may be undergoing early stages of apoptosis although this has not been conclusively proven. These cells may appear to contain MNs but should not be scored for MNs in the assay.
Karyorrhectic cells. These cells are characterized by the more extensive appearance of nuclear chromatin aggregation (relative to condensed chromatin cells) leading to fragmentation and eventual disintegration of the nucleus (Figure 3G). These cells may be undergoing a late stage of apoptosis but this has not been conclusively proven. These cells should not be scored for MNs in the assay.
Pyknotic cells. These cells are characterized by a small shrunken nucleus, with a high density of nuclear material that is uniformly but intensely stained (Figure 3H). The nuclear diameter is usually one- to two-thirds of a nucleus in normal differentiated cells. The precise biological significance of pyknotic cells is unknown but it is thought that these cells may be undergoing a form of cell death; however, the precise mechanism is unknown. These cells should not be scored for MNs in the assay.
Karyolytic cells. In these cells, the nucleus is completely depleted of DNA and apparent as a ghost-like image that has no Feulgen staining (Figure 3I). These cells thus appear to have no nucleus. It is probable that they represent a very late stage in the cell death process but this has not been conclusively proven. These cells should not be scored for MNs in the assay.
Scoring method
Initially 1000 cells were scored (500 per cytospin spot) per subject for all the various cells types outlined in the buccal cytome assay. These consisted of cells containing MNs, nuclear buds, basal cells, binucleates and the cell death parameters condensed chromatin, karyorrhectic, pyknotic and karyolytic cells. A total of 1000 differentiated and basal cells were scored in order to determine the frequency of MNs in a total of 1000 cells. Only basal and normal differentiated cells were scored for MNs and their scores were combined to give the overall incidence. Cells were scored using both bright field and fluorescence. Cells containing MNs on bright field were confirmed as being positive by examining the cells under fluorescence. The incidence of false positives can be minimized as DNA material such as nuclei and MNs fluoresce when viewed under fluorescence with a far-red filter.
Statistical analysis
One-way analysis of variance (ANOVA) was used to determine the significance of the cellular parameters measured between the older control, younger AD and older AD cohorts. Pairwise comparison of significance between these groups was determined using Tukey's test. ANOVA values, positive predictive values, negative predictive values, sensitivity, specificity, likelihood ratios and odds ratio were calculated using Graphpad PRISM (Graphpad Inc., San Diego, CA, USA). Cross-correlation analyses were performed using SPSS 14.0, SPSS Inc., Chicago. Significance was accepted at P < 0.05.
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Results |
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The results for DNA damage markers (cells with MNs or nuclear buds), cell proliferation markers (basal cells, binucleated cells) and cell death parameters (karyorrhectic cells, condensed chromatin cells, karyolytic cells and pyknotic cells) are summarized and illustrated in Figures 4–6. Cells with MNs or nuclear buds were slightly elevated in the younger AD group when compared to age-matched controls, although this difference did not achieve significance (P = 0.12 and P = 0.62, respectively). Micronucleated cells (88%) were in differentiated cells, whereas the remaining 12% were found in the basal cells. Basal cells were found to be lower by 81% in the younger AD and by 79% in the older AD cohorts when compared to the control group (P < 0.0001). Binucleate cell frequency showed no significant differences between the AD and control groups (P = 0.99). The frequency of cells with condensed chromatin was found to be reduced by 37.2% in the younger AD and by 56.6% in the older AD cohorts when compared to the control group (P < 0.0001). The frequency of karyorrhectic cells was found to be reduced by 82.9% in the younger AD and by 77.7% in the older AD cohorts when compared to the control group (P < 0.0001). Pyknotic and karyolytic cells were slightly lower in both the dementia groups but did not achieve significance when compared to the control (P = 0.47 and P = 0.84, respectively). There were no gender ratio differences between the control and the AD groups. There were no gender differences in biomarkers occurring within each cohort except for a significant increase in nuclear buds in males compared to females within the young AD group (P < 0.01). Cross-correlation analysis between MMSE scores and the biomarkers of the buccal cytome assay showed no positive correlation.
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Sensitivity and specificity
Values for both basal and karyorrhectic cells were analysed singularly and in combination to determine the sensitivity and specificity of these potential biomarkers which were most different between the control and AD groups. Table II lists the sensitivity and specificity data for diagnosis of AD based on basal and karyorrhectic cell frequency at different cut-off values. The odds ratio for being diagnosed with AD for a combined karyorrhectic and basal cell frequency value of <41 per 1000 cells is 140 with a specificity of 97% and a sensitivity of 82%. This would indicate that a false-positive rate for these potential diagnostic biomarkers within the general population would be 2.3% (positive predictive value of 97.7%) and that 23.1% (negative predictive value 76.9%) of those individuals tested that are likely to have AD would be falsely detected as normal. The relationship of basal cell and karyorrhectic cell frequencies shows a clear separation in distribution of these values for the control and AD cohorts (Figure 7).
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Discussion |
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This study is the first to use the buccal micronucleus cytome assay, which is based on cellular and nuclear morphology, to identify potential biomarkers that are associated with individuals that have just been clinically diagnosed with AD. Results showed a significant decrease in the number of basal cells (P < 0.0001), karyorrhectic cells (P < 0.0001) and condensed chromatin cells (P < 0.0001) in the AD patients, suggesting potential changes in the cellular kinetics or structural profile of the buccal mucosa. The combination of basal and karyorrhectic cells as potential biomarkers within the buccal mucosa may be useful as a possible future diagnostic to identify individuals with a high risk of developing or having AD.
The buccal mucosa is a stratified squamous epithelium consisting of four distinct layers (Figure 8). The stratum corneum or keratinized layer lines the oral cavity comprising cells that are constantly being lost as a result of everyday abrasive activities such as mastication. Below this layer lie the stratum granulosum or granular cell layer and the stratum spinosum or prickle cell layer containing populations of both differentiated and apoptotic cells. Integrated within these layers are convoluted structures known as rete pegs, containing the actively dividing basal cells known as the stratum germinativum which produce cells that differentiate and maintain the profile and integrity of the buccal mucosa.
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With increasing age the cell renewal process becomes less efficient and cellular regeneration decreases (25
,26). Ageing results in a decrease in the thickness of the epidermis and the underlying cell layers (27). Studies have shown that there is a decline in epidermal thickness with advancing age in both sexes with values occurring in the range from 11.4 to 22.6 µm (28). The number of cells within the stratum corneum does not diminish and therefore does not compromise its function as an essential protective layer (29). The rete pegs become less prominent and less convoluted resulting in a more flattened appearance (30,31). Further studies need to be performed, using histological sections and biopsies, to verify whether changes beyond normal ageing occur within the buccal mucosa of AD patients that could explain the reduced frequency of basal, condensed chromatin and karyorrhectic cells.
The results from the buccal cytome assay suggest significant changes within the BC profile of AD patients when compared to the control cohort. AD is histopathologically confirmed by the presence within the brain of neurofibrillary tangles consisting of the microtubule-associated protein tau and amyloid plaques which contain the 42 amino acid ß-amyloid (Aß42) resulting from the aberrant proteolysis of the amyloid precursor protein (APP). The gene for APP is found on chromosome 21 at 21q21 (32). It has been shown that both of these AD-associated proteins are present within vertebrate buccal mucosa (33,34). It has been shown that the level of tau in Alzheimer's oral epithelium had a significant positive correlation with tau levels in the cerebrospinal fluid (33). This indicates that BCs potentially may reflect some of the pathophysiological changes that occur within the AD brain. Individuals suffering from AD had significantly higher levels of oral epithelial tau when compared to controls who were clinically diagnosed as not suffering from AD (33). It has also been shown that Aß42 has been detected within the buccal mucosa of a genetically accelerated senescence mouse model (34) and that APP has been detected in the basal cell layer of human skin epidermis (35). APP at low levels (1–10 nM) has been shown to have a positive effect on the rate of proliferation of basal cells. However at higher concentrations of >10 nM, the rate of proliferation is significantly impaired. Whether increased tau, Aß42 or APP expression alters proliferation rate of the basal cells and cell ratios of buccal mucosa remains unclear. Only future studies combining these multiple parameters may answer this question.
Micronutrient deficiency such as zinc has also been shown to have a number of effects on the structural profile of the buccal mucosa (36,37). Rats fed on a zinc-deficient diet exhibited hyperplastic changes within the buccal mucosa. After 9 days, the keratin layer showed parakeratosis and significant thickening (37). After 18 days, all cells within the keratin layer showed further thickening and a complete parakeratotic transformation with a doubling of the mitotic rate within the basal cell layer. These changes were readily reversible once the rats returned to a normal control diet containing the normal amount of zinc (38). It is plausible that AD patients if suffering from a micronutrient deficiency such as zinc or if zinc is not bioavailable because it is aggregated by ß amyloid (39), then they may exhibit similar changes within the buccal mucosa. If such thickening were apparent, then the number of basal cells sampled would be significantly reduced as the keratinized layer thickened. Future studies need to be performed to investigate the effects of micronutrient zinc deficiency or availability of free zinc on changes in both cellular kinetics and structure of the AD buccal mucosa.
An alternative explanation for the reduced proportion of basal and apoptotic cells could be differences in cell sampling between AD and control groups. However, we think that this is unlikely because collections were done by one person following a standard protocol. Nevertheless, we performed studies with repeated sampling on the same day which showed that basal cells are increased as deeper layers are sampled (Table III). The proportion of karyorrhectic and condensed chromatin cells was found not to be significantly changed with repeated sampling (Table III). Although a trend for an increase of karyorrhectic cells with repeated sampling was evident (P-trend = 0.049), the increase in both basal cells and karyorrhectic cells at the fifth sample was 195 and 69% greater than the first sample. This is much less than the 427% increase in basal cells and 486% increase in karyorrhectic cells of the normal older controls relative to AD subjects. This argues against the possibility that observed differences between controls and AD may be explained by less vigorous sampling in AD compared to the controls. In addition, condensed chromatin cells were reduced in AD but remained unchanged with repeated sampling. More attention may be needed to standardize the sampling protocol in order to obtain reproducible results especially when multiple samplers are involved.
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Previous studies by Migliore and colleagues showed that micronucleus frequency is increased in lymphocytes and skin fibroblasts of AD patients (14
,16,18). However, our results in buccal mucosa do not show a marked difference between AD and controls for MNs frequency or nuclear buds. As expression of MNs requires nuclear division, MNs frequency in buccal mucosa is also dependent on the proportion of once-divided cells. It is possible that the reduced number of basal cells and perhaps a reduced proportion of dividing basal cells in AD may inhibit MNs and nuclear bud expression (the latter being S-phase dependent) (23). Our preliminary results need to be verified in larger cohorts and in other forms of dementia such as vascular dementia or Huntingdon's disease to determine whether the observed results are specific to AD patients. It would also need to be established, through further investigation, whether those individuals who have different genetic forms of AD reflect the same buccal cytome changes. If specificity for these buccal biomarkers could be verified for AD individuals, then further studies involving mild cognitively impaired individuals could be performed to determine if these changes occur in the early stages of AD. Buccal cytome changes may eventually be used to reflect disease severity or as a within-subject biomarker to gauge the effectiveness of preventative interventions aimed at slowing down the progression of the disease.
Changes in the ratios of BC populations may in the future form the basis of a non-invasive test that could identify potential biomarkers that are specific to individuals suffering from AD. Through further investigation, it may be possible to further refine the buccal cytome assay to include other DNA damage markers such as telomere shortening (40), aneuploidy (14,41) and oxidative stress (42,43) which have been shown to contribute to the etiology of AD. If combined with quantitative measures of tau and ß amyloid, a more comprehensive assay with potential diagnostic value could be developed and used to non-invasively identify presymptomatically individuals with increased risk of AD.
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Acknowledgments |
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The authors would like to thank Associate Professor Michael Roberts for his valuable contribution and comments during the compilation of the manuscript. The authors gratefully acknowledge the CSIRO clinic staff who aided in the sampling of the older control patients, especially Rosemary Macarthur and Carolyn Salisbury. Special thanks to Sue Moore for all her help and support during the course of the study. Lastly the authors greatly appreciate the efforts made by all the individuals who consented to participate in this study.
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Notes |
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* To whom correspondence should be addressed. Tel: +08 83038897; Fax: +08 83038899; Email: philip.thomas{at}csiro.au
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Received on May 16, 2007; revised on June 26, 2007; accepted on July 6, 2007.
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