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First published online 3 July 2008
doi: 10.1242/dev.020644
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1 Department of Neurobiology, 299 W. Campus Drive, Stanford University,
Stanford, CA 94305, USA.
2 Department of Pathology, Stanford University School of Medicine, Palo Alto, CA
94304, USA.
* Author for correspondence (e-mail: trc{at}stanford.edu)
Accepted 23 May 2008
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SUMMARY |
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 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key words: Drosophila, ROS, Degeneration, Mitochondria, Succinate dehydrogenase, Synapse
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INTRODUCTION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). By
contrast, mutations that directly affect mitochondrial metabolism cause
early-onset neurodegeneration in a number of developmental disorders,
including Leigh Syndrome, Leber's hereditary optic neuropathy and Kearns-Sayre
Syndrome (reviewed by Vogel,
2001). In Leigh Syndrome, for example, patients are apparently
normal at birth, but become broadly ataxic, display nystagmus and spasticity
over the first few months of life, and ultimately die during childhood
(Leigh, 1951). These
behavioral deficits are associated with lesions in several sub-cortical
structures, as well as optic atrophy
(Leigh, 1951). However, no
animal model that recapitulates the neurodegenerative changes associated with
these early-onset diseases has been described, and the molecular mechanisms
that would link mitochondrial dysfunction to degeneration are incompletely
understood. Here, we establish an animal model of Leigh Syndrome in
Drosophila photoreceptors.
The fly visual system is a powerful model for understanding
neurodevelopment, and for defining the mechanisms that underlie human
neurodegenerative diseases (reviewed by
Marsh and Thompson, 2006;
Clandinin and Zipursky, 2002).
In particular, the synaptic connections between photoreceptor axon terminals
and their post-synaptic targets have been described using both light and
electron microscopy, and many molecular components of the synapse have been
identified (Prokop and Meinertzhagen,
2006; Hiesinger et al.,
2005; Zinsmaier et al.,
1994). In addition, Drosophila photoreceptors faithfully
recapitulate cellular pathologies associated with Parkinson's disease and
polyglutamine repeat diseases like Huntington's disease, and have provided a
powerful platform for examining the genetic interactions that influence
disease progression (Greene et al.,
2003; Jackson et al.,
1998). Finally, the Drosophila retina has also been used
to examine the molecular mechanisms that regulate mitochondrial trafficking
and activity, and thus provides a wealth of reagents for examining
mitochondrial function (Stowers et al.,
2002; Gorska-Andrzejak et al.,
2003; Mandal et al.,
2005). Here, we demonstrate that mild disruption of mitochondrial
function is sufficient to induce degeneration in Drosophila
photoreceptors, and we define the molecular mechanisms necessary and
sufficient for synapse loss in this context.
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MATERIALS AND METHODS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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;
Grigliatti, 1986;
Clandinin et al., 2001).
Optomotor assays were conducted as previously described
(Clandinin et al., 2001). To
generate flies in which retinas were almost completely homozygous for
SdhA, while the rest of the fly was heterozygous, we used the FLP/FRT
system, expressing the FLP recombinase under the control of two different
eyeless promoters, paired with a recessive cell lethal
(Newsome et al., 2000;
Chotard et al., 2005). A
similar strategy, replacing the cell-lethal chromosome with one bearing GFP
under the control of the eye-specific GMR promoter
(Chang et al., 2002), was used
to generate negatively marked clones. To generate retinas both homozygous for
SdhA and expressing CuZnSOD, we incorporated an Act5a FRT STOP FRT
CuZnSOD flip-out transgene (Sun and Tower,
1999; Sun et al.,
2002). To generate retinas homozygous for both SdhA and
Miro, we generated stocks that contained both mutations on FRT
chromosomes (and a corresponding control stock), a stock that contained two
appropriate cell lethals, and a FLP source (under the control of the eyeless
promoter). The following mutants and transgenes were used:
SdhA1110 and SdhA1404 (both from this
work); Sdh5 and Sdh7 (P. Lawrence,
University of Cambridge, UK); Mirosd32 and UAS-mitoGFP (T.
L. Schwarz, Harvard Medical School, Boston, MA); and UAS-Buffy (N. Bonini,
University of Pennsylvania, Philadelphia, PA).
|
). We
visualized all photoreceptor (R) cells using the monoclonal antibody mAb24B10
chaoptin at 1:50, synaptic vesicles using mAb1G12 Cysteine
String Protein at 1:10 and active zones with mAbnc82 at 1:50 [all from the
Developmental Studies Hybridoma Bank (DSHB) at the University of Iowa]. We
visualized mitochondria with the monoclonal antibody MS507 Complex V at
1:500 (Mitosciences). Third instar larval eye discs were assessed in whole
mount using mAb24B10 (1:50) and Bar (1:100)
(Higashijima et al., 1992).
Third instar optic lobe development was assessed using mAb24B10 (1:50), Rat
elaV (DSHB; 1:100), Goat HRP-FITC (1:100; Jackson
ImmunoResearch) and mouse Repo (1:100; DSHB). Secondary antibodies were
obtained from Invitrogen. Fluorescence images were collected on a Leica TCS
SP2 AOBS confocal microscope, visualized using Imaris (Bitplane), and mounted
using Adobe Photoshop. Quantification of CSP staining was performed by
selecting the entire lamina as a region of interest in a single section,
thresholding the signal, and measuring the fraction of the region [ImageJ
(NIH)]. TUNEL staining was performed as described
(Bilen et al., 2006). Staining
of activated Drice was performed as described
(Yoo et al., 2002). SDH
activity was assessed in whole-mount third instar larval eye discs made
homozygous for either SdhA or a control chromosome. Eye discs were
stained by using the activity assay described by Pearse
(Pearse, 1972). Sections from
adult fly retinas were prepared as described
(Sullivan et al., 2000).
Transmission electron microscopic analysis was performed as described, except
that heads were mounted in Epon (Pesah et
al., 2004). Counts were made from three flies of each genotype,
with 10-15 R cell terminals being analyzed per fly.
ATP assay
Retinas from somatic mosaic adult flies were dissected on ice, and assayed
using a luciferin/luciferase-based ATP assay kit (Calbiochem). This dissection
tears the retina along the fenestrated membrane at its base, and includes all
of the R cell cell bodies, but excludes all brain tissue from the tissue
preparation. In our hands, by using the FLP recombinase and cell-lethal
combination, less than 1% of retinal tissue is not rendered homozygous. To
normalize for differences in the amount of retinal tissue in each experimental
sample, the amount of pigment in each specimen was measured at 280 nm using a
spectrophotometer (Pharmacia), and compared against a standard curve
containing different amounts of retinal tissue. The signal was corrected for
non-specific absorbance by retinal tissue by subtracting the observed
absorbance from that seen in the equivalent number of white mutant
retinas. To determine the amount of ATP measured in each sample, we generated
a standard curve using known quantities of ATP. To calculate the cellular
concentration of ATP, we directly determined the mass of a large quantity of
dissected adult retinas and calculated the volume based on an estimated
specific gravity of 1.05. This measured volume was essentially identical to
the volume of the retina, calculated based on its physical dimensions and
geometry (data not shown).
Antioxidant treatment
Flies were raised on standard fly food treated with either 200 µg/ml
alpha-tocopherol in ethanol, or ethanol alone. Adult flies were transferred to
freshly treated food on the day of eclosion.
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RESULTS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). To identify genes involved in the formation and maintenance
of this structure, we conducted a forward genetic screen based on a behavioral
assay that depended on the ability of adult flies to respond to motion cues
(Clandinin et al., 2001). To
identify genes whose loss-of-function phenotypes would otherwise be lethal, we
conducted this screen in somatic mosaic animals in which only photoreceptor
cells were made homozygous mutant, while the rest of the animal is
heterozygous (Stowers et al., 1999;
Newsome et al., 2000). By
using this approach, we identified two mutations in the succinate
dehydrogenase flavoprotein subunit (SdhA, CG17246) of mitochondrial
complex II, that we designated SdhA1110 and
SdhA1404. Two additional alleles,
SdhA5 and SdhA7 were identified in
separate forward genetic screen for the loss of complex II activity
(Lawrence, 1981). All four
alleles were recessive lethal, with a lethal phase extending to the first
larval stage. All four alleles failed to complement one another and a small
deficiency, uncovering nine genes for this phenotype. Finally, all
trans-heterozygote combinations, both bearing the deficiency and bearing the
reference allele SdhA1110, had lethal phases in late
embryogenesis and the first larval stage (see Table S1 in the supplementary
material). These genetic observations demonstrate that all four alleles are
strong reduction-of-function mutations.
The succinate dehydrogenase complex comprises four subunits, catalyzes the
oxidation of succinate to fumarate, and transfers electrons into the electron
transport chain (ETC) via ubiquinone (Fig.
1A). This complex is not essential for respiration, as electrons
can also enter the ETC via complex I and can be passed directly to complex
III, bypassing complex II. Thus, disrupting this complex should allow
oxidative phosphorylation to continue. The flavoprotein subunit,
SdhA, contains two domains, a FAD-binding 2 domain, which covalently
binds a flavin adenine dinucleotide (FAD) cofactor, and the succinate
dehydrogenase flavoprotein C-terminal domain, which forms the catalytic site
of the enzyme (Fig. 1B)
(Yankovskaya et al., 2003).
This protein is highly conserved between flies and humans
(Fig. 1B). DNA sequence
analysis revealed a single missense mutation in each of our SdhA
alleles that caused a non-conservative change in an amino acid residue in SdhA
(Fig. 1B). These changes lie in
both the FAD-binding 2 domain (SdhA1110,
SdhA1404 and SdhA7) and the C-terminal
domain (SdhA5).
|
). To directly
compare the enzymatic functions of complex II in wild-type and mutant retinas,
we generated mitotic clones by expressing the yeast FLP recombinase under the
control of the eyeless promoter, and examined developing eye discs of
third instar larvae homozygous for either a control chromosome
(Fig. 1C) or
SdhA1110 (Fig.
1D). In wild-type eye discs, SDH activity was relatively low in
retinal precursor cells, but increased rapidly in differentiating
photoreceptor cells, and outlined developing ommatidial pre-clusters
(Fig. 1C). By contrast, the
activity of SDH was strongly reduced in retinas homozygous for
SdhA1110. Similar results were obtained with
SdhA1404 (data not shown), and had previously been
described in other imaginal discs using SdhA5 and
SdhA7 (Lawrence,
1981). To confirm that this activity corresponded to the function
of complex II, we added malonate, which competes with succinate at the
catalytic site of the holoenzyme. The addition of this inhibitor strongly
reduced the amount of activity detected, and eliminated all differences in
staining intensity across the disc (Fig.
1E). Thus, this assay captures qualitatively the activity of
complex II, and confirms the genetic observations, indicating that
SdhA1110 and SdhA1404 alleles are
strong reduction-of-function alleles of SdhA.
SdhA mutant R cells form normal synaptic terminals, which progressively degenerate
Adult eyes in which all R cells are homozygous for SdhA mutations
are outwardly indistinguishable from wild-type controls (data not shown). To
examine the underlying neural structures, we first visualized R cell
projections into the first optic ganglion, the lamina, using whole-eye clones
(see Fig. S2 in the supplementary material). By late pupal development (72
hours after puparium formation, APF), R cells have formed a regular array of
axon fascicles, termed cartridges, each comprising six photoreceptor axon
terminals surrounding their post-synaptic targets, the lamina neurons (see
Fig. S1 in the supplementary material). At this stage, the terminals of
control R cells, as well as those of R cells homozygous mutant for
SdhA, expressed high levels of the synaptic vesicle component
Cysteine String Protein 2A, as well as of the active zone marker Bruchpilot
(Zinsmaier et al., 1994;
Wagh et al., 2006). In mutant,
but not control, flies these structures degenerated progressively over the
first 5 days after eclosion (Fig.
2; see Fig. S2 in the supplementary material). This phenotype was
maintained when we generated SdhA clones by expressing FLP
recombinase under the control of a different retina-specific promoter,
ey3.5, was independent of which cell-lethal mutation was
used to increase clone size, and was seen in all four SdhA alleles
(data not shown). Thus R cells mutant for SdhA form normal synaptic
terminals during development, as assessed using confocal microscopy, which
degenerate later in the late pupal and adult fly.
To observe these changes more closely, we made small, negatively marked, SdhA mutant clones. In wild-type R cells on the day of eclosion, cartridge structure and the localization of synaptic vesicles was highly regular (Fig. 3A,B). This expression pattern was maintained for at least the first 5 days of adult life (Fig. 3E,F). On the day of eclosion, R cell terminals mutant for SdhA appeared normal (Fig. 3C,D). However, as we had observed in the whole-eye clones, by 5 days after eclosion, R cell terminals lacked almost all labeling of synaptic vesicles (Fig. 3G,H). Thus, these small clones recapitulate the phenotype observed in large clones, albeit with a transient difference in phenotypic onset observed at eclosion, which is likely to be a result of increased perdurance.
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), and retinas homozygous for SdhA mutations were
indistinguishable from controls (Fig.
4A,D). We next sectioned adult retinas on the day of eclosion, as
well as 5 days later. In control flies, ommatidia form a regular array of
facets composed of R cells and surrounding pigment cells, which can be easily
visualized during the first 5 days of adulthood
(Fig. 4B,C). However, in
SdhA mutant animals the retina was disrupted at eclosion: although
apparently normal rhabdomes were still visible, R cell bodies were variable in
size, and the array of ommatidia was disordered
(Fig. 4E). Five days after
eclosion, this phenotype was more severe: many cell bodies were swollen, and
many had lost their rhabdomes (Fig.
4F). We conclude that mutant photoreceptor structures in the
retina form normally and then progressively degenerate in the adult fly.
Moreover, because SdhA mutant retinas were of normal morphology and
pigmentation, it appears that this disruption affects R cells more severely
than the non-neuronal pigment cells (which were also homozygous for
SdhA). To test whether these phenotypes specifically reflect late-stage degeneration, we systematically examined the early stages of photoreceptor differentiation and visual system development in animals in which photoreceptors were homozygous mutant for SdhA (see Fig. S3 in the supplementary material). In particular, during the third larval stage, we demonstrated that SdhA mutant R cells assembled into ommatidia normally, and expressed fate-appropriate markers. Moreover, the targeting of SdhA mutant R cell axons to appropriate ganglia, the development of their target neurons in the lamina, and the recruitment of their associated glia all occurred normally. Finally, by injecting fluorescent dye into single ommatidia during mid-pupal development, we demonstrated that SdhA mutant R cells almost invariably chose the appropriate post-synaptic partners. Thus, loss of SdhA function in R cells causes little, if any, phenotype, until at least the very late stages of pupal development.
Mitochondrial density and morphology are aberrant in SdhA mutant R cell terminals
We next examined whether functional deficits in complex II activity were
associated with changes in mitochondrial localization within R cell terminals
(Fig. 5). To do this, we
labeled mitochondria either by expressing mitochondrially localized GFP
(mitoGFP) under the control of rhodopsin1 GAL4 (specific to R1-R6 cells), or
by staining with an antibody directed against the alpha subunit of the
mitochondrial ATP synthase, complex V. In control animals, mitochondria were
enriched in R cell synaptic terminals during adulthood
(Fig. 5B,C,J,K). By comparison,
in SdhA mutant R cells, the intensity of mitochondrial staining was
reduced at eclosion (Fig.
5F,G), and continued to decline such that, by 5 days after
eclosion, there was little or no mitochondrial staining within photoreceptor
terminals (Fig. 5N,O).
|
). Moreover R cell terminals contained numerous
mitochondrial profiles (Fig.
6A,A') (Stowers et al.,
2002). By contrast, in SdhA mutant animals, some R cell
terminals lacked mitochondrial profiles
(Fig. 6B,B'), while
others contained small, abnormal mitochondria
(Fig. 6C,C'). As a
result, the mean ratio of the volume of mitochondrial profiles per R cell
terminal volume in SdhA mutants was 0.121±0.018, compared with
0.224±0.015 in control flies (P<0.001;
Fig. 6F). We note that the
average number of mitochondria per terminal did not change in SdhA
mutants (data not shown). Furthermore, whereas mitochondria in wild-type
terminals frequently displayed the characteristic invaginated cristae
(Fig. 6A,D), mitchondria in
SdhA mutant R cell terminals were frequently abnormal in morphology,
displaying irregular invaginations (Fig.
6C). Characteristic of neural degeneration, we also observed
multi-lamellar bodies (Fig.
6B). Five days after eclosion, lamina neuropil containing
wild-type R cell terminals retained their structure and organization
(Fig. 6D,D'), whereas
those containing SdhA mutant R cell terminals had severely
degenerated (Fig. 6E,E').
Cells containing capitate projections were not observed, so R cell terminals
could not be identified (Fig.
6E,E'). Thus, these ultrastructural studies reveal that
SdhA mutations cause defects in mitochondrial volume and structure
consistent with our observations of reduced complex V staining in flies at the
day of eclosion.
Cells mutant for SdhA are not depleted of ATP
We reasoned that disruption of the ETC in SdhA mutant R cells
could impair oxidative phosphorylation sufficiently to cause ATP depletion,
which might, in turn, lead to synapse loss and degeneration. To test this
possibility, we measured ATP levels in control and mutant retinas using a
biochemical assay, and observed no difference in the average amount of ATP per
retina, both 0 days and 5 days after eclosion
(Fig.7A). To normalize for
differences in the mass of retina isolated in each experimental sample, we
measured the screening pigment absorbance in each specimen using a
spectrophotometer, and then compared each measurement to a standard curve
generated using known quantities of retina. We found no difference in
absorbance between control and SdhA mutant retinas. This standard
curve was corrected for non-specific absorbance by retinal tissue by
subtracting the observed absorbance from that seen in the equivalent number of
white mutant retinas. The amount of pigmentation in wild-type and
SdhA mutant retinas was equivalent (see Fig. S4 in the supplementary
material). By directly measuring the mass of isolated retinas, and by using a
standard curve generated with known quantities of ATP, we then calculated that
the cellular concentration of ATP was between 1.5 mM and 1.8 mM. As nearly a
third of the mass of the retina is acellular [comprising the lenses of each
ommatidium, and the vitreous humor
(Franceschini and Kirschfeld,
1971)], these values reflect a conservative estimate of the
concentration of ATP within R cells. Thus, SdhA mutant retinas
contain normal levels of ATP in the cell soma. These results are consistent
with the fact that R cell differentiation and development in SdhA
mutants was normal, as previous observations had demonstrated that
mitochondrial mutations that significantly reduce ATP levels (by 40%) cause
dramatic developmental defects (Mandal et
al., 2005).
|
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). As
dark-rearing has no effect on the development of R cells and their synapses in
wild-type flies (Fig.
7B,D,F,H), we examined whether raising SdhA mutant
animals in complete darkness could suppress photoreceptor degeneration.
However, by comparison with flies raised under standard 12:12 light:dark
conditions, dark-reared flies showed indistinguishable levels of synaptic
degeneration in the brain (Fig.
7C,G) and damage in the retina
(Fig. 7E,I). Thus, we conclude
that the degenerative phenotypes we observe in SdhA mutant animals do
not result from ATP depletion.
|
).
Notably, impairment of the ETC increases ROS formation
(Staniek and Nohl, 2000). We
reasoned that if ROS contributes to the degeneration we observe, prolonged
treatment with the antioxidant alpha-tocopherol should suppress this
phenotype. To test this, we grew control and SdhA mutant somatic
mosaic flies on standard food supplemented with 200 µg/ml alpha-tocopherol.
On the day of eclosion, flies were placed on newly treated food, and then
examined 5 days later. Chronic treatment with alpha-tocopherol had no affect
on the development of the lamina or retina in control flies
(Fig. 8A,F,K), and treatment
with vehicle alone did not suppress the degeneration observed in SdhA
mutants (Fig. 8B,G,L). However,
treatment with alpha-tocopherol strongly suppressed the loss of synaptic
vesicles (Fig. 2B,
Fig. 8C,D,H,I), as well as the
loss of mitochondrial staining and the disorder of the active zone (see Fig.
S5 in the supplementary material), in SdhA1110 and
SdhA1404 mutant R cells at 5 days after eclosion.
Intriguingly, antioxidant treatment failed to suppress the swelling of
photoreceptor cell bodies and the degeneration of rhabdomes
(Fig. 2A,
Fig. 8M,N).
To exclude the possibility that these protective effects of
alpha-tocopherol might reflect a protective function of this molecule that is
independent of its effect on ROS levels, we exploited a parallel genetic
approach to reducing ROS. Superoxide dismutase (SOD) is a central component of
the cellular defense against ROS, and acts by converting superoxide to
less-reactive metabolites (McCord et al., 1969). We therefore tested whether
overexpression of SOD in R cells could suppress the degeneration we observed
in SdhA mutants. To overexpress this enzyme specifically in the
retina, we generated eye-specific mosaic flies mutant for SdhA, and
used the yeast FLP/FRT system to induce eye-specific expression of CuZnSOD
under the control of the actin promoter, using the Act5a FRT STOP FRT CuZnSOD
flip-out transgene (Sun and Tower,
1999; Sun et al.,
2002). As in other control flies, overexpression of the SOD
transgene in otherwise wild-type R cells did not disrupt R cell morphology in
the retina, or the lamina (data not shown). However, consistent with the
notion that ROS are crucial mediators of synapse loss in SdhA mutant
R cells, CuZnSOD overexpression dramatically increased the level of synaptic
vesicle staining in many R cell terminals through 5 days after eclosion
(Fig. 2B,
Fig. 8E,J). Consistent with the
effect we observed when SdhA mutants were treated with antioxidants,
this suppression was uncoupled from effects on the retina: CuZnSOD expression
had no effect on retinal degeneration (Fig.
2A, Fig. 8O).
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SdhA mutant mitochondria act in the cell body to induce synaptic degeneration
ROS are highly reactive, and hence act locally with respect to their site
of production. We therefore sought to determine whether the synaptic
degeneration observed in SdhA mutant R cells reflects damage caused
by mitochondria located locally in the synaptic terminal, or could be caused
by damage elsewhere in the cell. To distinguish between these alternatives, we
excluded mitochondria from the synaptic terminal using the mitochondrial
trafficking mutant Miro. When the mitochondrial trafficking pathway
controlled by Miro is blocked, mitochondria remain restricted to the cell body
beginning in the early stages of R cell axonal differentiation and never enter
the brain; during adulthood mitochondria in these mutants are at least 20
µm from the terminal (Stowers et al.,
2002; Gorska-Andrzejak et al.,
2003). We reasoned that if SdhA mutant mitochondria acted
locally to damage the synaptic terminal, preventing mitochondria from entering
the terminal should suppress degeneration. If, however, damage induced by
mitochondria elsewhere in the cell is sufficient to cause degeneration, then
preventing entry should have no effect. Using markers for R cell terminal
morphology and synapse formation, we first determined that Miro
mutant R cell terminals were indistinguishable from controls at eclosion, and
did not degenerate like SdhA mutant photoreceptors
(Fig. 9A-C,E-G,I-K). Although
the presence of mitochondria at the synaptic terminal is necessary for the
normal formation of synapses at the neuromuscular junction
(Guo et al., 2005), our
results are consistent with previous studies that demonstrated that
synapse-associated mitochondria are not required for synapse development in R
cells (Stowers et al., 2002).
However, R cells doubly homozygous for mutations in SdhA and
Miro displayed a degeneration phenotype indistinguishable from that
seen in SdhA single mutant cells
(Fig. 2B,
Fig. 9D,H,L). Thus, removal of
abnormal mitochondria from the terminal is not sufficient to prevent
degeneration, demonstrating that ROS-induced damage in the cell body can cause
the synapse loss we observe in SdhA mutants. Moreover, these
experiments demonstrate blocking mitochondrial trafficking into the synaptic
terminal is not sufficient to cause synapse degeneration, and thus the ROS
produced in SdhA mutant photoreceptors cannot simply block
mitochondrial transport.
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DISCUSSION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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A fly model of Leigh Syndrome
Our work describes the first animal model of Leigh Syndrome that
recapitulates the neurodegenerative changes seen in human patients. Leigh
Syndrome is caused by inherited mutations in proteins crucial to electron
transport, including mutations in SDH subunits
(Bourgeron et al., 1995;
Horvath et al., 2006).
Children born with such mutations appear normal during the first months of
life, but then show signs of psychomotor delay and regression prior to death.
These deficits are associated with widespread degeneration in many
sub-cortical structures (Leigh,
1951). Notably, optic atrophy is also frequently observed
(Birch-Machin et al., 2000;
Leigh, 1951). In many
respects, our findings in the fly parallel this time course: R cells mutant
for SdhA develop completely normally, adopt the correct cell fates,
innervate the appropriate synaptic partners, and assemble synapses normally.
However, beginning around the time of eclosion, R cells degenerate,
progressively losing expression of synaptic markers, and undergoing extensive
morphological changes. Thus, our model captures many elements of the human
disease.
Genetic analysis of the succinate dehydrogenase complex
Mutations in other SDH subunits have been described in worms and in flies.
The complex is composed of four subunits: the flavoprotein subunit (SdhA) and
an iron-sulfur subunit (SdhB) that together make up the catalytic core of the
holoenzyme; and two membrane-bound subunits (SdhC and SdhD), which anchor the
complex to the inner mitochondrial membrane and transfer electrons to
ubiquinone (Ackrell et al.,
1990). Mutations in SdhB in Drosophila
(Walker et al., 2006), and in
SdhC (mev-1) in C. elegans
(Ishii et al., 1998), shorten
life span in a high oxygen environment. In the fly, this sensitivity to
hyperoxia manifests as morphological abnormalities in the mitochondria of
flight muscles, and behavioral deficits in geotaxis
(Walker et al., 2006). In
worms, mutations in SdhC increase ROS production under normal oxygen
tension (Senoo-Matsuda et al.,
2001). These previous findings are consistent with our results in
that they link deficits in mitochondrial complex II activity to the production
of excess ROS. However, unlike our mutations in SdhA, neither of
these mutations are homozygous lethal, suggesting that they cause
comparatively weaker effects on the activity of the complex. Moreover, in
neither case was gene function examined in the context of degenerative changes
in specific neurons. Finally, the degenerative phenotypes we observed are
distinct from those associated with blocking mitochondrial protein translation
(Chihara et al., 2007). That
is, although SdhA mutant R cells display degeneration of axons,
specifically blocking mitochondrial translation in olfactory projection
neurons causes only degeneration of dendrites.
What are the targets of ROS?
Given the causal role for ROS in neurodegeneration, identifying the
cellular site of action of ROS represents a crucial challenge. One model is
that the primary effect of excessive ROS production is to cause damage to
mitochondria, forming a positive-feedback loop in which ROS-induced damage
further enhances ROS production (reviewed by
Lin and Beal, 2006). Indeed,
in our system, the damage we observed to synaptic mitochondria in
SdhA mutant R cells is ROS dependent, as it can be blocked by
antioxidant treatment. However, damage to mitochondria may not be the direct
cause of synapse loss, because synaptic degeneration could be suppressed by
overexpression of CuZnSOD, a form of SOD that is thought to act primarily in
the cytoplasm (O'Brien et al.,
2004). That is, although damage to mitochondria is clearly an
important aspect of the cellular degeneration we see, excess ROS appears to
directly alter the activities of one or more cytosolic components to cause
synapse loss. Our observations also demonstrate that degenerative changes in
the cell body can be uncoupled from those in the synaptic terminal, as
overexpression of CuZnSOD, or addition of an exogenous antioxidant, suppressed
the SdhA mutant phenotype in the brain without mitigating the retinal
phenotype. This differential sensitivity could reflect quantitative
differences in the ability of one structure to withstand damage more than
another, or could reflect qualitative differences in the specific cellular
targets affected by ROS. One possibility, then, is that mitochondrial
dysfunction activates two molecularly distinct pathways, one that is mediated
by excess ROS and causes synapse loss, and one that is mediated by
as-yet-unknown components that leads to the degeneration of components of the
cell body. Finally, our demonstration that removal of SdhA mutant
mitochondria from the synaptic terminal is not sufficient to prevent synaptic
degeneration suggests that the critical cellular targets of ROS that affect
synapse structure are in the cell body, not the terminal.
A common mechanism of neurodegeneration?
The mechanisms we describe here are likely to underlie important aspects of
other neurodegenerative disease pathologies. In particular, alterations in
mitochondrial complex II activity have also been linked to Huntington's
disease (HD) (reviewed by Walker,
2007). A specific decrease in the expression of two SDH subunits,
SDHA and SDHB, occurs in striatal neurons of Huntington's patients, and
ectopic expression of mutant Huntingtin causes a similar decrease in neuronal
cultures (Benchoua et al.,
2006). Moreover, overexpression of these SDH subunits suppresses
the cell death induced by mutant Huntingtin protein both in cultured striatal
neurons, and in yeast (Benchoua et al.,
2006; Solans et al.,
2006). In this yeast model of HD, reductions in SDH activity are
also associated with increased ROS production
(Solans et al., 2006).
Finally, chronic administration of 3-nitropropionic acid, an inhibitor of SDH,
to rodents and primates recapitulates many of the neurological deficits seen
in HD patients (Palfi et al.,
1996; Guyot et al.,
1997; Brouillet et al.,
1998). As our studies demonstrate that mutations in SdhA
are sufficient to cause neurodegeneration through increased ROS production, we
speculate that, in striatal neurons, this constitutes one of the central
mechanisms underlying neurodegeneration seen in HD. More broadly, our studies
are consistent with the notion that the excessive production of ROS that has
been detected in other neurodegenerative diseases, such as Parkinson's disease
and ALS, might provide a sufficient explanation for at least some of the
neurodegenerative changes seen in these disorders.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/135/15/2669/DC1
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ACKNOWLEDGMENTS |
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REFERENCES |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Ackrell, B. A. C., Johnson, M. K., Gunsalus, R. P. and Cecchini, G. (1990). Structure and function of succinate dehydrogenase and fumarate reductase. In Chemistry and Biochemistry of Flavoproteins. Vol. 2 (ed. F. Muller), pp.229 -297. Boca Raton, FL: CRC Press.
Ashburner, M. (1989). Drosophila: a Laboratory Manual. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press.
Balaban, R. S., Nemoto, S. and Finkel, T. (2005). Mitochondria, oxidants, and aging. Cell 120,483 -495.[CrossRef][Medline]
Benchoua, A., Trioulier, Y., Zala, D., Gaillard, M. C., Lefort,
N., Dufour, N., Saudou, F., Elalouf, J. M., Hirsch, E., Hantraye, P. et
al. (2006). Involvement of mitochondrial complex II defects
in neuronal death produced by N-terminus fragment of mutated huntingtin.
Mol. Biol. Cell 17,1652
-1663.
Bilen, J., Liu, N., Burnett, B., Pittman, R. and Bonini, N. (2006). MicroRNA Pathways Modulate Polyglutamine-Induced Neurodegeneration. Mol. Cell 24,157 -163.[CrossRef][Medline]
Birch-Machin, M. A., Taylor, R. W., Cochran, B., Ackrell, B. A. and Turnbull, D. M. (2000). Late-onset optic atrophy, ataxia, and myopathy associated with a mutation of a complex II gene. Ann. Neurol. 48,330 -335.[CrossRef][Medline]
Bourgeron, T., Rustin, P., Chretien, D., Birch-Machin, M., Bourgeois, M., Viegas-Pequignot, E., Munnich, A. and Rotig, A. (1995). Mutation of a nuclear succinate dehydrogenase gene results in mitochondrial respiratory chain deficiency. Nat. Genet. 11,144 -149.[CrossRef][Medline]
Brouillet, E., Guyot, M. C., Mittoux, V., Altairac, S., Conde, F., Palfi, S. and Hantraye, P. (1998). Partial inhibition of brain succinate dehydrogenase by 3-nitropropionic acid is sufficient to initiate striatal degeneration in rat. J. Neurochem. 70,794 -805.[Medline]
Chang, H. C., Newmyer, S. L., Hull, M. J., Ebersold, M., Schmid,
S. L. and Mellman, I. (2002). Hsc70 is required for
endocytosis and clathrin function in Drosophila. J. Cell
Biol. 159,477
-487.
Chihara, T., Luginbuhl, D. and Luo, L. (2007). Cytoplasmic and mitochondrial protein translation in axonal and dendritic terminal arborization. Nat. Neurosci. 10,828 -837.[CrossRef][Medline]
Chotard, C., Leung, W. and Salecker, I. (2005). glial cells missing and gcm2 cell autonomously regulate both glial and neuronal development in the visual system of Drosophila. Neuron 48,237 -251.[CrossRef][Medline]
Clandinin, T. R. and Zipursky, S. L. (2002). Making connections in the fly visual system. Neuron 35,827 -841.[CrossRef][Medline]
Clandinin, T. R., Lee, C. H., Herman, T., Lee, R. C., Yang, A. Y., Ovasapyan, S. and Zipursky, S. L. (2001). Drosophila LAR regulates R1-R6 and R7 target specificity in the visual system. Neuron 32,237 -248.[CrossRef][Medline]
Franceschini, N. and Kirschfeld, K. (1971). Pseudopupil phenomena in the compound eye of Drosophila. Kybernetik 9,159 -182.[CrossRef][Medline]
Gorska-Andrzejak, J., Stowers, R. S., Borycz, J., Kostyleva, R., Schwarz, T. L. and Meinertzhagen, I. A. (2003). Mitochondria are redistributed in Drosophila photoreceptors lacking milton, a kinesin-associated protein. J. Comp. Neurol. 463,372 -388.[CrossRef][Medline]
Greene, J. C., Whitworth, A. J., Kuo, I., Andrews, L. A., Feany,
M. B. and Pallanck, L. J. (2003). Mitochondrial pathology and
apoptotic muscle degeneration in Drosophila parkin mutants. Proc.
Natl. Acad. Sci. USA 100,4078
-4083.
Grigliatti, T. (1986). Mutagenesis in Drosophila: A Practical Approach (ed. D.B. Roberts), pp.39 -48. Oxford: IRL Press.
Guo, X., Macleod, G. T., Wellington, A., Hu, F., Panchumarthi, S., Schoenfield, M., Marin, L., Charlton, M. P., Atwood, H. L. and Zinsmaier, K. E. (2005). The GTPase dMiro is required for axonal transport of mitochondria to Drosophila synapses. Neuron 47,379 -393.[CrossRef][Medline]
Guyot, M. C., Hantraye, P., Dolan, R., Palfi, S., Maziére, M. and Brouillet, E. (1997). Quantifiable Bradykinesia, gait abnormalities, and Huntington's disease-like striatal lesions in rats chronically treated with 3-nitropropionic acid. Neuroscience 79,45 -56.[CrossRef][Medline]
Hiesinger, P. R., Fayyazuddin, A., Mehta, S. Q., Rosenmund, T., Schulze, K. L., Zhai, R. G., Verstreken, P., Cao, Y., Zhou, Y., Kunz, J. et al. (2005). The v-ATPase V0 subunit a1 is required for a late step in synaptic vesicle exocytosis in Drosophila. Cell 121,607 -620.[CrossRef][Medline]
Higashijima, S., Michiue, T., Emori, Y. and Saigo, K.
(1992). Subtype determination of Drosophila embryonic external
sensory organs by redundant homeo box genes BarH1 and BarH2. Genes
Dev. 6,1005
-1018.
Horvath, R., Abicht, A., Holinski-Feder, E., Laner, A., Gempel,
K., Prokisch, H., Lochmuller, H., Klopstock, T. and Jaksch, M.
(2006). Leigh syndrome caused by mutations in the flavoprotein
(Fp) subunit of succinate dehydrogenase (SDHA). J. Neurol.
Neurosurg. Psychiatr. 77,74
-76.
Ishii, N., Fujii, M., Hartman, P. S., Tsuda, M., Yasuda, K., Senoo-Matsuda, N., Yanase, S., Ayusawa, D. and Suzuki, K. (1998). A mutation in succinate dehydrogenase cytochrome b causes oxidative stress and ageing in nematodes. Nature 394,694 -697.[CrossRef][Medline]
Jackson, G. R., Salecker, I., Dong, X., Yao, X., Arnheim, N., Faber, P. W., MacDonald, M. E. and Zipursky, S. L. (1998). Polyglutamine-expanded human huntingtin transgenes induce degeneration of Drosophila photoreceptor neurons. Neuron 21,633 -642.[CrossRef][Medline]
Lawrence, P. A. (1981). A general cell marker for clonal analysis of Drosophila development. J. Embryol. Exp. Morphol. 64,321 -332.[Medline]
Leigh, D. (1951). Subacute necrotizing
ecephalomyelopathy in an infant. J. Neurol. Neurosurg.
Psychiatr. 14,216
-221.
Lin, M. T. and Beal, M. F. (2006). Mitochondrial dysfunction and oxidative stress in neurodegenerative diseases. Nature 443,787 -795.[CrossRef][Medline]
Mandal, S., Guptan, P., Owusu-Ansah, E. and Banerjee, U. (2005). Mitochondrial regulation of cell cycle progression during development as revealed by the tenured mutation in Drosophila. Dev. Cell 9,843 -854.[CrossRef][Medline]
Marsh, J. L. and Thompson, L. M. (2006). Drosophila in the study of neurodegenerative disease. Neuron 52,169 -178.[Medline]
McCord, J. M. and Fridovich, I. (1969).
Superoxide dismutase. An enzymic function for erythrocuprein (hemocuprein).
J. Biol. Chem. 244,6049
-6055.
Meinertzhagen, I. A. and Hanson, T. E. (1993). The Development of Drosophila Melanogaster (ed. M. Bate and A. Martinez Arias). Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press.
Newsome, T. P., Asling, B. and Dickson, B. J. (2000). Analysis of Drosophila photoreceptor axon guidance in eye-specific mosaics. Development 127,851 -860.[Abstract]
Niven, J. E., Anderson, J. C. and Laughlin, S. B. (2007). Fly photoreceptors demonstrate Energy-Information trade-offs in neural coding. PLoS Biol. 5, 828-840.
O'Brien, K. M., Dirmeier, R., Engle, M. and Poyton, R. O.
(2004). Mitochondrial protein oxidation in yeast mutants lacking
manganese-(MnSOD) or copper- and zinc-containing superoxide dismutase
(CuZnSOD): evidence that MnSOD and CuZnSOD have both unique and overlapping
functions in protecting mitochondrial proteins from oxidative damage.
J. Biol. Chem. 279,51817
-51827.
Palfi, S., Ferrante, R. J., Brouillet, E., Beal, M. F., Dolan,
R., Guyot, M. C., Peschanski, M. and Hantraye, P. (1996).
Chronic 3-nitropropionic acid treatment in baboons replicates the cognitive
and motor deficits of Huntington's disease. J.
Neurosci. 16,3019
-3025.
Pearse, A. G. E. (1972). Histochemistry. Vol. 2, 3rd edn. London: Churchill Livingston.
Pesah, Y., Pham, T., Burgess, H., Middlebrooks, B., Verstreken,
P., Zhou, Y., Harding, M., Bellen, H. and Mardon, G. (2004).
Drosophila parkin mutants have decreased mass and cell size and increased
sensitivity to oxygen radical stress. Development
131,2183
-2194.
Prokop, A. and Meinertzhagen, I. A. (2006). Development and structure of synaptic contacts in Drosophila. Semin. Cell Dev. Biol. 1, 20-30.
Senoo-Matsuda, N., Yasuda, K., Tsuda, M., Ohkubo, T., Yoshimura,
S., Nakazawa, H., Hartman, P. S. and Ishii, N. (2001). A
defect in the cytochrome b large subunit in complex II causes both superoxide
anion overproduction and abnormal energy metabolism in Caenorhabditis elegans.
J. Biol. Chem. 276,41553
-41558.
Solans, A., Zambrano, A., Rodriguez, M. and Barrientos, A.
(2006). Cytotoxicity of a mutant huntingtin fragment in yeast
involves early alterations in mitochondrial OXPHOS complexes II and III.
Hum. Mol. Genet. 15,3063
-3081.
Staniek, K. and Nohl, H. (2000). Are mitochondria a permanent source of reactive oxygen species? Biochim. Biophys. Acta 1460,268 -275.[Medline]
Stowers, R. S. and Schwarz, T. L. (1999). A
genetic method for generating Drosophila eyes composed exclusively of mitotic
clones of a single genotype. Genetics
152,1631
-1639.
Stowers, R. S., Megeath, L. J., Gorska-Andrzejak, J., Meinertzhagen, I. A. and Schwarz, T. L. (2002). Axonal transport of mitochondria to synapses depends on milton, a novel Drosophila protein. Neuron 36,1063 -1077.[CrossRef][Medline]
Sullivan, W., Ashburner, M. and Hawley, R. S. (2000). Drosophila Protocols. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press.
Sun, J and Tower, J. (1999). FLP
recombinase-mediated induction of Cu/Zn-superoxide dismutase transgene
expression can extend the life span of adult Drosophila melanogaster flies.
Mol. Cell. Biol. 19,216
-228.
Sun, J., Folk, D., Bradley, T. J. and Tower, J.
(2002). Induced overexpression of mitochondrial Mn-superoxide
dismutase extends the life span of adult Drosophila melanogaster.
Genetics 161,661
-672.
Vogel, H. (2001). Mitochondrial myopathies and the role of the pathologist in the molecular era. J. Neuropathol. Exp. Neurol. 60,217 -227.[Medline]
Wagh, D. A., Rasse, T. M., Asan, E., Hofbauer, A., Schwenkert, I., Durrbeck, H., Buchner, S., Dabauvalle, M. C., Schmidt, M., Qin, G. et al. (2006). Bruchpilot, a protein with homology to ELKS/CAST, is required for structural integrity and function of synaptic active zones in Drosophila. Neuron 49,833 -844.[CrossRef][Medline]
Walker, F. O. (2007). Huntington's disease. Lancet 369,218 -228.[CrossRef][Medline]
Walker, D. W., Hájek, P., Muffat, J., Knoepfle, D.,
Cornelison, S., Attardi, G. and Benzer, S. (2006).
Hypersensitivity to oxygen and shortened lifespan in a Drosophila
mitochondrial complex II mutant. Proc. Natl. Acad. Sci.
USA 103,16382
-16387.
Yankovskaya, V., Horsefield, R., Törnroth, S., Luna-Chavez,
C., Miyoshi, H., Léger, C., Byrne, B., Cecchini, G. and Iwata, S.
(2003). Architecture of succinate dehydrogenase and reactive
oxygen species generation. Science
299,700
-704.
Yoo, S. J., Huh, J. R., Muro, I., Yu, H., Wang, L., Wang, S. L., Feldman, R. M., Clem, R. J., Müller, H. A. and Hay, B. A. (2002). Hid, Rpr and Grim negatively regulate DIAP1 levels through distinct mechanisms. Nat. Cell Biol. 4, 416-424.[CrossRef][Medline]
Zinsmaier, K. E., Eberle, K. K., Buchner, E., Walter, N. and
Benzer, S. (1994). Paralysis and early death in cysteine
string protein mutants of Drosophila. Science
263,977
-980.
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