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From the Kinsmen Laboratory of Neurological Research, University of British Columbia, Vancouver, British Columbia, Canada
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Studies of the pathology of Alzheimer's disease (AD) have done much to focus attention on the larger role of the complement system in health and disease and particularly to highlight the role of endogenous brain cells, including neurons, in inflammatory and immune responses. It has long been known that activated components of the classical complement pathway are associated with AD lesions.1-5 Initially, it was believed that such activation was secondary to a classical antigen-antibody reaction in AD brain. However, failure to confirm the presence of antibodies inspired the search for alternative activators of complement. ß-Amyloid protein (Aß), the donor peptide of AD amyloid deposits, was found to bind C1q and activate complement in vitro.6 Other molecules associated with AD lesions which activate complement in vitro are amyloid P, C-reactive protein, and Hageman factor (reviewed in Ref. 5 ). Thus, the complement cascade could be activated in vivo by one or more of these molecules. The possibility that brain itself acts as the source of complement proteins was also investigated, as a serum source was unlikely due to the blood-brain barrier. Astrocytes, microglia, and neuroblastoma cells were all found to produce complement proteins in vitro.7–11 In vivo immunohistochemical12,13 and in situ hybridization analyses14,15 have indicated that neurons are the most abundant source of brain complement proteins, at least in the hippocampus and temporal cortex.
The plaque and tangle lesions of AD are chronic, and the association of complement proteins with them could reflect a low level process of little pathological significance, with most of the deposits reflecting events remote in time. On the other hand, they could reflect an aggressive level of activity which could be an important contributor to AD pathogenesis.
To assess the relative level of complement activity in AD compared with control cases, we used the reverse transcriptase-polymerase chain reaction (RT-PCR) technique to measure relative levels of the mRNAs for all of the components of the classical complement pathway, ie, C1q, C1r, C1s, C2, C3, C4, C5, C6, C7, C8, and C9. We assayed 11 regions of brain, as well as liver, heart, kidney and spleen of many of these cases. All tissues studied contained detectable levels of most of the complement protein mRNAs. The levels in brain were strikingly elevated in AD compared with controls, particularly in those areas with heavy pathological involvement. However, levels in the liver and other organs were not affected. Western blot analyses showed that the complement mRNAs were translated into their protein products, and that, in AD brain, the classical complement pathway was strongly activated, including assembly of the membrane attack complex. These data provide evidence of the generation of complement mRNAs in AD brain, with continuous activation of the classical complement pathway. Such activity points to the probable role of complement as a major force driving pathology in the disease.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Fifteen autopsy cases were used in this study. Brain, heart,
liver, spleen, and kidney were investigated, but not all organs were
obtained from every case. Details are given in Table 1
. The brains from four cases of AD and
five without neurological disease were studied. In each case of AD, the
clinical diagnosis was confirmed by routine pathological analysis for
plaques and tangles. All cases met standard criteria for moderate to
severe AD. No AD pathology was noted in the control brains. Peripheral
organs only were obtained from an additional six cases. The age range
of the AD cases was 65 to 78 years (70.5 ± 2.8), with postmortem
delays varying from 6 to 16 hours. The ages of the control brain cases
ranged from 43 to 82 years (72.2 ± 7.3), with postmortem delays
from 9 to 48 hours. Postmortem delay was not considered to be a
contributing factor to the results since we have previously established
that there is almost no observable degradation for the mRNAs of
cyclophilin and the complement proteins in tissue stored for periods of
up to 6 days in the cold.16
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The techniques used for RNA extraction and preparation of RT-PCR products have been described in detail elsewhere.16 Briefly, total RNA was extracted from approximately 500 mg of each tissue sample by the acid guanidinium thiocyanate-phenol-chloroform method. The samples were treated with 10 U of RNase-free DNase (Pharmacia) for 60 minutes at 37°C in 25 µl of 1x reverse transcriptase buffer (50 mmol/L Tris-HCl, 75 mmol/L KCl, 3 mmol/L MgCl2) containing 40 U of RNase inhibitor (Pharmacia) and 1 mmol/L dithiothreitol, followed by an incubation at 85°C for 5 minutes to inactivate the enzyme.
Single-strand cDNA synthesis was performed on 5 µg of total RNA extract. The reaction mixture contained the RNA sample, 25 µl of 1x reverse transcriptase buffer containing 1 µg of random hexamer primers (pDN6, Pharmacia), 1 mmol/L deoxynucleotides (GIBCO BRL), 5 mmol/L dithiothreitol, 40 U of RNase inhibitor (Pharmacia) and 500 U of reverse transcriptase (Superscript TMII RT, GIBCO BRL). Duplicate assays were carried out at 42°C for 90 minutes, followed by heat inactivation at 65°C for 10 minutes.
The appropriate quantity of cDNA was covered with 50 µl of mineral oil and amplified in a 50-µl reaction buffer containing 67 mmol/L Tris-HCl (pH 8.8), 16.6 mmol/L ammonium sulfate, 10 mmol/L 2-mercaptoethanol, 200 mmol/L dNTPs, 2 mmol/L MgCl2, 40 pmol of each specific oligonucleotide primer, and 2.5 U of Taq DNA polymerase (GIBCO BRL). The thermal profile used on a Fisher Scientific (Toronto, Canada) programmable thermal controller consisted of a denaturation step of 94°C for 1 minute, an annealing step of 55°C for 30 seconds, and an extension step of 72°C for 1 minute. The extension step in the first cycle was for 3 minutes at 72°C. All samples were initially denatured for a total of 5 minutes (94°C).
To determine appropriate parameters for amplification of the cDNA products, the method of Nakayama et al17 was followed. The method involves using graded amounts of cDNA and varying cycles of amplification to determine a range where the logarithm of reaction product intensity is linear with the cycle number. In preliminary experiments, we tested amounts of cDNA from 0.01 µg to 1 µg and cycle numbers from 20 to 40, using the standard conditions described above. We found that for cyclophilin the log of PCR product intensity was linearly proportional to cycle number from 20 to 29 cycles, and, for all complement genes, from 31 to 37 cycles. Plateaus were reached after 29 and 37 cycles, respectively. The product intensity was proportional to the cDNA concentration from 0.1 µg to 1 µg. Accordingly, the standard PCR procedure adopted was 0.5 µg of cDNA (1 µl) and 27 cycles of amplification for cyclophilin and 35 cycles for the complement products.
Each PCR reaction product was electrophoresed through a 6%
polyacrylamide gel and the product visualized by incubation for 10
minutes in a solution containing 10 ng/ml of ethidium bromide.
Resulting gel bands were imaged using a GDS 6700 image analyzer (Ultra
Violet Products, Uplands, CA). The relative intensities of the bands,
expressed as optical density units, were quantitatively analyzed using
NIH Image software 1.61. Each complement mRNA analysis was made in
parallel with a cyclophilin mRNA analysis to provide an internal
standard. Cyclophilin has been widely used as a gene product with
constant expression in tissue including brain.18
Polaroid
photographs of the gels were taken. Table 2
lists the GenBank accession number for
the gene sequences used and the positions chosen for primer design.
Except for C1r, they spanned 1 to 3 introns. The primer sequences were
as previously published.16
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lists the restriction enzymes used and the products anticipated
from the known GenBank sequences. In every case, an initial PCR product
of the correct size was obtained, and restriction enzyme products of
the correct size were obtained (Table 2)
. Western blots were performed on extracts of the soluble fraction of homogenates of control and AD hippocampus and compared with normal human serum and and serum activated by aggregated IgG. Brain samples were homogenized in 5x vol/protein extraction buffer (0.02 mol/L Tris-HCl, pH 7.5) containing the protease inhibitors phenylmethylsulfonyl fluoride (1 µg/ml) and aprotinin (1 µg/ml), and 1 mmol/L EDTA. Homogenates were centrifuged at 18,000 x g at 4°C for 30 minutes. The protein content of the supernatants was determined,19 and the samples were diluted in SDS sample buffer (60 mmol/L Tris, pH 6.8; 2.5% SDS, 5% ß-mercaptoethanol) to a final protein content of 1 µg/ml and were boiled for 3 minutes. Samples containing 20 µg of protein were loaded onto 7.5% acrylamide minigels.
Normal human serum taken from a 44-year-old male volunteer was diluted 1:20 in Veronal buffer. A 2-ml aliquot of the diluted serum was mixed with 50 ml of a solution of 2 µg/ml heat-aggregated human IgG (Sigma). The mixture was incubated at 37°C for 1 hour. Aliquots of the normal and IgG activated serum were then diluted in 2 volumes of SDS buffer and boiled for 3 minutes. Samples containing 20 µg of protein were loaded onto 7.5% acrylamide minigels. Life Technologies high range prestained standards were used as molecular weight markers. After 45 minutes of electrophoresis (200 V), the proteins were transferred onto nitrocellulose membranes (Immobilon P, Millipore Corp, MA) at 7 V for 48 minutes using a semidry blotter.
Due to the high molecular weight of the membrane attack complex, modifications of the electrophoresis and protein transfer steps were required. A 3% polyacrylamide gel was used, and separation was carried out for 2.5 hours at 100 V in a cold room with the apparatus surrounded by ice. Transfer to membranes was then carried out at 100 V for 5 hours in the cold.
Membranes were blocked in 5% skim milk for 2 hours. The immunoblots
were next treated for 2 hours at room temperature with a primary
anti-complement antibody, followed by treatment for 1 hour with an
appropriate secondary antibody labeled with horseradish peroxidase. The
primary and secondary antibodies are listed in Table 3
. Immunoreactivity was visualized by
incubation with Supersignal CL-HRP chemiluminescent substrate (Pierce
Chemical Co., Rockford, IL). After draining, the membranes were covered
in clear plastic wrapping and exposed to x-ray film (Hyper Film ECL,
Amersham Life Sciences) for 0.3 to 2 minutes, depending on the strength
of the signal.
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Immunohistochemistry was performed as previously
reported.13
Briefly, 30-µm sections were cut on a
freezing microtome. Free-floating sections were first treated for 30
minutes with 0.3% H2O2 solution in 0.01 mol/L
phosphate-buffered saline, pH 7.4, containing 0.3% Triton X-100 to
reduce endogenous peroxidase activity. They were then incubated
overnight at 4°C with one of the primary antibodies shown in Table 3
.
The sections were washed and treated with appropriate biotinylated
secondary antibodies (Table 3)
for 2 hours at room temperature,
followed by incubation in avidin-biotinylated horseradish peroxidase
complex (ABC Elite, Vector) for 1 hour at room temperature. Peroxidase
labeling was visualized by incubation of the sections in 0.01%
3,3'-diaminobenzidine (Sigma) containing 0.6% nickel ammonium sulfate
and 0.00015% H2O2 in 0.05 mol/L Tris-HCl
buffer, pH 7.6. When a dark purple color developed, sections were
washed, mounted on glass slides, and coverslipped with Entellan.
Controls were performed by omitting the primary antibody.
Statistical Analysis
Data are expressed as means ± SE. The data were analyzed by
analysis of variance, followed by Student t-tests, for the
significance of the difference between AD and control across all
complement mRNAs and regions, and for the individual complement mRNA by
region. Correlation analyses were used to determine whether there was
any significant relationship between the level of each brain mRNA
studied and postmortem delay in either the control or AD group. The
Holm multiple comparison method20
was applied to each set
of analyses. Significance was accepted at P < 0.05.
Data were analyzed as directly obtained and also as values normalized
to the cyclophilin internal standard in the same sample. Cyclophilin
levels typically fell within a range of 1% (Tables 2 and 4)
, so the normalization adjustment was
minor and did not affect the statistical differences found.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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is an
example of an ethidium bromide-stained gel showing the individual
products for each of the complement mRNAs. Single products were
obtained of the predicted size. The cDNA of each product was purified
and subjected to selective endonuclease digestion. In each case, the
predicted number of products of the expected size was obtained. Details
are shown in Table 2
.
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Figure 2
shows bar graphs in which
average normalized complement mRNA levels of AD and control cases are
compared for each of the 11 areas of brain studied. Notice in all of
the bar graphs how closely comparable the cyclophilin values were for
normal and AD cases. As can be seen from the graphs, C3 and C4 mRNAs
were the most highly expressed of all of the complement mRNAs in all
areas of brain in both AD and normal cases. In control brain, C1q and
C9 mRNAs were expressed at the lowest levels, with the other complement
mRNAs lying in between. The expression of all complement mRNAs
varied little from area to area in control brains although C8 mRNA
was not detected in the occipital and motor cortices.
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). In the
cerebellum, which is only mildly affected in AD, C1q mRNA increased
from 1.39 in controls to only 3.30 in AD cases. C9 mRNA increased from
0.65 in controls to 6.98 in AD (see graphs in Figure 2
). Other
complement mRNAs followed the same general pattern of up-regulation in
AD, although the up-regulation was not as great as that observed for
C1q and C9 mRNAs. The least up-regulation was for C3 and
C4 mRNAs, which were expressed at the highest levels. The other
areas of brain, which were intermediate in their pathological
involvement, were also intermediate in their mRNA up-regulation.
Complement mRNAs were also measured in the liver, heart, spleen, and
kidney of AD and control cases. The results are shown in Table 4
.
Inspection of this table shows that the mRNA levels in these peripheral
areas were highly comparable between AD and control cases, with no
statistically significant differences between them being detected. It
is noteworthy, however, that the complement mRNA levels in affected
areas of AD brain were substantially higher than those in AD or control
liver and other peripheral organs.
Figure 3
shows the results of Western
blot experiments of normal serum (lane 1), of extracts of control
hippocampus (lane 2) and AD hippocampus (lane 3), and of the same serum
as lane 1 with aggregated IgG added to activate complement (lane 4).
Strong bands were detected for all of the complement proteins in the AD
hippocampal extract and IgG treated serum, including bands for the
activation fragments C3d, C4d, and the membrane attack complex. These
same bands were detected to a much lesser extent in the normal serum
sample, possibly indicating mild in vivo activation. By
contrast, bands were not detected in the control hippocampal extract
for C1q, C1r, C1s, C2, C5, C6, C7, C8, C9, and the membrane attack
complex. The complement protein fragments represented by these bands
have previously been reported for Western blots of heart
extracts.16
These data indicate that the complement
mRNAs detected in RT-PCR experiments are being translated
into their protein products, and that the classical pathway is
being activated in tissue.
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illustrates by
immunohistochemistry association of some of the complement proteins
with the lesions of AD. Antibodies to C1q, the component which
identifies the target tissue and initiates the complement cascade,
stain senile plaques prominently, as well as tangled pyramidal neurons
in the AD transentorhinal cortex (Figure 4A)
. Control transentorhinal
cortex, under the same conditions, shows no immunostaining (Figure 4B)
.
Very intense staining of senile plaques in the same area in AD is found
for C4d, a fragment characteristic of classical pathway activation
(Figure 4C)
. Again, no staining is seen in control tissue (Figure 4D)
.
The antibody to the membrane attack complex prominently stains
dystrophic neurites and intracellular tangles in AD
transentorhinal cortex (Figure 4E)
, while no staining is obtained in
control tissue (Figure 4F)
. Since the membrane attack complex can
destroy host cells through the process of bystander lysis, the evidence
of binding to dystrophic neurites in Figure 4E
is suggestive of
autodestruction of tissue.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Here we have made semiquantitative comparisons of the mRNA levels of all components of the classical complement pathway, ie, C1q, C1r, C1s, C2, C3, C4, C5, C6, C7, C8, and C9, showing that they are easily detected in all 11 areas of AD brain studied. They are also detectable in normal brain tissue, with the minor exception of C8 in the occipital and motor cortices. C3 and C4 were the most abundantly expressed of the mRNAs in all areas of both normal and AD tissue. They showed little variation from area to area in normal tissue, and showed less than a twofold up-regulation in AD brain tissue, even in the most heavily affected areas. In contrast, C1q, the component which activates the pathway by binding to target tissue, and C9, the component which gives functional capacity to the membrane attack complex, were expressed at the lowest levels in normal tissue but showed dramatic up-regulation in affected AD brain areas.
The mRNA levels for the housekeeping gene cyclophilin were remarkably constant in all areas and for all cases, with values generally falling within 1% of each other and with an outside range of less than 4%. Cyclophilin was also unaffected by postmortem delay, as we have previously reported.16 The constancy of cyclophilin mRNA levels indicates that the methodology itself is highly reproducible and that the gene product is unaffected by the various pathologies and postmortem delay. The complement mRNAs were similarly unaffected by postmortem delay in normal and AD brains. The high variability in complement mRNA values observed between highly affected and mildly affected areas of AD brain is therefore probably reflective of the physiological state of the tissue and not other extraneous factors.
All of the complement mRNAs were, as expected, strongly detected in
liver of AD and non-AD cases. The levels in AD and non-AD liver were
highly comparable (Table 4)
. By contrast, the levels in AD hippocampus
were substantially higher for all of the complement components than in
AD or control liver (Table 4)
. These data are again consistent with the
concept of high local production of complement mRNAs in affected areas
of AD brain, with continuous activation of the classical pathway.
The mRNAs for all complement components were also detected in spleen,
kidney, and heart (Table 4)
. In general, these levels were comparable
to those observed in normal brain, with that for C1q being the lowest
and those for C3 and C4 the highest. The fact that all organs produce
mRNAs for all complement proteins suggests that complement production
is far more ubiquitous than is generally recognized and that local
production and activation may be associated with a wide spectrum of
pathological disorders.
We have previously reported on the expression of complement mRNAs in isolated rabbit hearts and their up-regulation after reperfusion.25 We have also reported on the presence of complement mRNAs and their protein products in human heart and their up-regulation after myocardial infarcts.16 Activated complement fragments were detected on damaged myocardium of previously as well as recently infarcted tissue. This indicates that complement-mediated damage can continue to accumulate for years after an initial insult, and, in the case of heart, this insult may be as mild as an anoxic episode.
The previously reported association of activated complement proteins with AD lesions does not indicate when, and to what degree, the complement pathway has been activated. The process could have been proceeding at a low level for years. However, the high up-regulation of mRNAs, and the appearance of strong bands in Western blots for complement activation products, indicates that vigorous activation of the pathway must be continuously taking place. Because the complement system powerfully drives inflammation and tissue destruction in a highly targeted manner, these results imply a substantial contribution of complement to AD pathology.
The process by which the endogenously produced complement proteins are activated is not known with certainty. However, the disease process in AD involves continuous deposition of amyloid deposits. These deposits contain the in vitro activators ß-amyloid protein,6 amyloid P,26-29 C-reactive protein,30 and Hageman factor,31 which should provide a strong stimulus for in vivo activation. Extracellular neurofibrillary tangles, which are also being continuously formed as neurons with intracellular tangles die, have deposited on them the complement activator amyloid P.32 The high up-regulation of mRNAs in areas where plaques and tangles accumulate may explain why these areas are highly vulnerable to the AD pathological process.
There are now considerable data suggesting that inflammatory processes drive the pathology in AD. Multiple epidemiological studies show that individuals taking anti-inflammatory drugs or suffering from conditions where such drugs are routinely administered, have a substantially reduced prevalence of AD.33,34 Moreover, a small double-blind, placebo-controlled clinical trial of the anti-inflammatory drug indomethacin demonstrated an apparent arrest of the disease process.35 Complement may set the pace of neuronal degeneration. If this is true, complement inhibitors might prove to be highly effective therapeutic agents in AD.
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Acknowledgements |
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Footnotes |
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Supported by a grant from the Jack Brown and Family A.D. Research Fund and donations from individual British Columbians.
Accepted for publication December 22, 1998.
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