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First published online 30 August 2006
doi: 10.1242/dev.02575
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Department of Molecular and Cellular Biology, Life Sciences South Room 531, University of Arizona, Tucson, AZ 85721, USA.
* Author for correspondence (e-mail: fares{at}email.arizona.edu)
Accepted 8 August 2006
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SUMMARY |
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 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key words: C. elegans, CUP-5, Mucolipidosis Type IV, ABC Transporter, MRP-4
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INTRODUCTION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Bach,
2001). The first class, which includes corneal clouding and
achlorhydria is due to the accumulation of large lipid-rich vacuoles in the
respective tissues. The second class, which includes psychomotor retardation,
is due to neuronal cell death. The gene responsible for this disease encodes
the protein human mucolipin 1, which belongs to the `transient receptor
potential' family of proteins (Bargal et
al., 2000; Bassi et al.,
2000; Sun et al.,
2000). Human mucolipin 1 functions as a non-selective cation
channel, the activity of which is modulated by pH
(LaPlante et al., 2002;
Raychowdhury et al.,
2004).
The C. elegans CUP-5 protein is the ortholog of human mucolipin 1
(Fares and Greenwald, 2001b).
Mutations in cup-5 result in the appearance of large vacuoles and the
absence of lysosomal degradation in various cell types because of a defect in
lysosome biogenesis (Fares and Greenwald,
2001b; Hersh et al.,
2002; Treusch et al.,
2004). The large vacuoles are hybrids of late endosomes and
lysosomes and represent the terminal endocytic compartments, at least in
scavenger cells called coelomocytes
(Treusch et al., 2004). In
cup-5 null embryos, the degradation of yolk and of membrane proteins
is significantly delayed, particularly in developing intestinal cells
(Schaheen et al., 2006). As in
other tissues, this defect is accompanied by an expansion in the size of the
terminal compartment. Furthermore, this leads to ectopic apoptosis, to
developmental abnormalities and ultimately to organism death primarily because
of the developmental defects (Hersh et
al., 2002; Schaheen et al.,
2006).
We describe in this study the identification and the characterization of a
suppressor of the lysosomal defect and the lethality of cup-5(null)
worms. The suppressor, MRP-4, is a member of the ATP-binding cassette (ABC)
transporter superfamily that is widely distributed in prokaryotes and
eukaryotes (Bauer et al., 1999;
Cheong et al., 2006;
Dean et al., 2001;
Holland and Holland, 2005;
Sheps et al., 2004). ABC
transporters bind ATP and use the energy to transport various molecules,
including drugs, toxins, peptides and lipid derivatives across the membrane of
various cellular compartments. This family of proteins contain several
transmembrane helices and ATP-binding domains typically containing three
conserved Walker domains, and have been implicated in various physiological
processes, including pleiotropic drug and heavy metal resistance and membrane
trafficking (Dean et al.,
2001; Holland and Holland,
2005).
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MATERIALS AND METHODS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). Transgenic
integrated lines were made by bombardment of unc-119(ed3) worms using
pHD137 as a co-bombardment marker (Praitis
et al., 2001). RNAi was done by the feeding method as previously
described (Timmons and Fire,
1998). Markers used were: pgp-2(gk114) I
(Nunes et al., 2005);
cup-5(ar465) III (Fares and
Greenwald, 2001b); cup-5(zu223) III
(Hersh et al., 2002);
unc-36(e251) III (closely linked to cup-5)
(Brenner, 1974;
Fares and Greenwald, 2001a;
Fares and Greenwald, 2001b);
unc-69(e587) III (Brenner,
1974); ced-9(n1950n2161) III
(Gumienny et al., 1999);
rme-2((b1008) IV (Grant and
Hirsh, 1999); cpl-1(ok360) V
(Britton and Murray, 2004);
rme-1(b1045) V (Grant et al.,
2001); cup-10(ar479) V
(Dang et al., 2003);
rme-6(b1018) X (Sato et al.,
2005); bIs1[VIT-2::GFP; pRF4] (which expresses a fusion
of the worm YP170 protein VIT-2 to GFP)
(Grant and Hirsh, 1999);
cdIs77[GFP::LGG-1; unc-119(+)-pmyo-2::GFP] X (which
expresses GFP-LGG-1 in all cells and GFP in the pharynx)
(Schaheen et al., 2006);
nT1[qIs51] IV, V (is a dominant balancer)
(Siegfried et al., 2004); and
qC1 (a balancer chromosome that suppresses recombination between
cup-5 and unc-36)
(Graham and Kimble, 1993).
cup-5(zu223) unc-36(e251) worms bearing various transgenes were
isolated from qC1-balanced parent heterozygotes; the eggs from these
homozygous progeny were analyzed in the various assays. unc-69(e587)
ced-9(n1950n2161) worms were isolated from qC1-balanced parent
heterozygotes; the eggs from these homozygous progeny were analyzed.
cpl-1(ok360) worms were identified as progeny that did not express
GFP in the pharynx from cpl-1(ok360)/nt1[qIs51] heterozygotes; the
eggs from these homozygous progeny were analyzed.
Identification of mrp-4(cd8)
We mutagenized cup-5(zu223) unc-36(e251)/qC1; arIs37
hermaphrodites with EMS as previously described
(Brenner, 1974). We then
picked 239 F1 wild-type progeny (predicted to be heterozygous for new
mutations) to separate plates. From each of these 239 plates, we picked 12 Unc
F2 progeny and screened for viable F3 Unc progeny. These were backcrossed four
times to wild-type worms to confirm the suppression and to remove background
mutations. We identified the single cd8 allele in this screen.
We mapped cd8 between stp156 and stp72 on
chromosome X using sequence-tagged site (sTs) mapping, as previously described
(Fares and Greenwald, 2001a;
Williams et al., 1992). To
determine the actual mutated gene, we used an RNAi screen of all the genes on
chromosome X that would rescue the embryonic lethality of cup-5(zu223)
unc-36(e251). We only identified one gene, mrp4, that mapped
between stp156 and stp72, and that phenocopied the
suppression by the cd8 allele. Sequencing of the open reading frame
in cd8 worms confirmed the presence of a mutation. Furthermore, we
could reverse the suppression of mrp-4(cd8); cup-5(zu223)
unc-36(e251) by expressing wild type mrp-4 from a transgenic
array.
Methylpyruvate feeding
Methylpyruvate (Sigma, St Louis, MO) was added to the NGM medium at a
concentration of 14 mM before the plates were poured. In addition, OP50 was
spun down and resuspended in a 14 mM solution of methylpyruvate before seeding
the plates.
TUNEL assay
The TUNEL assay was carried out as previously described
(Schaheen et al., 2006).
LysoTracker red staining
LysoTracker Red (Invitrogen, Carlsbad, CA) was added to the NGM medium at a
concentration of 50 nM before the plates were poured. In addition, OP50 was
spun down and resuspended in a 50 nM solution of LysoTracker Red before
seeding the plates.
Nile Red staining
Worms and eggs were incubated in a 0.06 mg/ml solution of Nile Red
(Invitrogen, Carlsbad, CA) for 1 hour. These were then washed once with M9 and
once with 1 mM levamisole before imaging.
Measuring viability
Adult worms were allowed to lay eggs overnight at 20°C. The adults were
then removed and the percentage of eggs that hatched after one day was
determined at 20°C. The percentage of hatched eggs that developed to
adults was determined after 2 more days at 20°C. Each experiment was
repeated five times and at least 90 eggs were counted in each case. The
results represent the average of these experiments and the bars represent
standard deviations.
Measurements and statistical methods
To determine sizes of compartments, confocal images of embryos were opened
using Adobe Photoshop (Adobe Systems, San Jose, CA) and analyzed without any
modification. At least 50 discrete intracellular structures from several
embryos were selected and the number of pixels included was determined. The
reported values reflect the average surface areas of the highlighted
structures and the standard deviations (1 pixel is approximately 0.01
µm2).
To quantitate intensity of MRP-4::GFP or Nile Red staining, confocal images of embryos were opened using ImageJ software (National Institutes of Health, Bethesda, MD) and analyzed without any modification. At least 50 discrete intracellular structures from several embryos were selected and the integrated density (mean intensity divided by area) was determined. The reported values reflect the average integrated densities of the highlighted structures and the standard deviations.
Student's t-test was used to compare average measurements from two samples using a two-tailed distribution (Tails=2) and a two-sample unequal variance (Type=2).
The bars on the charts represent standard deviations.
cDNA sequencing
We sequenced the mrp-4 cDNA clone yk86a11. We also used 5'
RACE to identify the 5' end of the mrp-4 mRNA (Roche,
Indianapolis, IN). These results confirmed the predicted splicing pattern of
mrp-4.
Molecular methods
Standard methods were used for the manipulation of recombinant DNA
(Sambrook et al., 1989).
Polymerase chain reaction (PCR) was done using the Expand Long Template PCR
System (Boehringer Manheim, Manheim, Germany), according to the manufacturer's
instructions. All other enzymes were from New England Biolabs (Beverly, MA)
unless otherwise indicated. The transcriptional fusion of the mrp-4
promoter to GFP was made by PCR amplifying 3 kb of genomic sequences upstream
of F21G4.2 using the forward primer (5'
CACACAGCATGCCACACATCTATTAGGAATGTG 3') and the reverse primer (5'
CACACAACCGGTCCGTATCTTCTCTCCTTATTTCG 3'). The resulting PCR product was
digested with SphI and AgeI and subcloned into the same
sites of pPD95.75. The MRP-4::GFP translational fusion was made by PCR
amplifying 8.1 kb of genomic sequences using the forward primer (5'
CACACAGCATGCCATTCGTATATATCCTCC 3') and the reverse primer (5'
CACACAACCGGTATGAGGCCAGCGCGTTTGGCCATTGAGTAG 3'). The resulting PCR
product was digested with SphI and AgeI and subcloned into
the same sites of pPD95.75. This results in GFP fused to the C terminus of
MRP-4 under the control of 2.7 kb of genomic sequences upstream of the F21G4.2
start codon. The plasmid pHD137 carries both wild type unc-119 and
pmyo-2::GFP and was made by subcloning the 2.7 kb
SphI-ApaI fragment from pPD118.33 into the
XhoI-ApaI sites of plasmid MM106B after blunt-ending with T4
DNA Polymerase (Praitis et al.,
2001).
Microscopy
Adult worms were allowed to lay eggs for one day on the NGM plates,
sometimes supplemented with LysoTracker Red. Embryos were placed in M9 buffer
for imaging. Confocal images were taken with a Nikon PCM 2000, using HeNe 543
nm excitation for the red dye and argon 488 nm for the green dye. Exactly the
same exposure and magnification was used to capture the embryos from different
strains that expressed the same marker.
Immunofluorescence staining of embryos using the IBF-2 antibody MH33 were
done as previously described (Bossinger et
al., 2004). Immunofluorescence staining of embryos using
antibodies against RME-2 was done as previously described
(Britton and Murray, 2004).
Immunofluorescence staining of embryos using the LIN-12 antibody was done as
previously described (Hermann et al.,
2000). Exactly the same exposure and magnification was used to
capture the embryos from different strains stained with the same
antibodies.
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RESULTS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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A phylogenetic analysis of the 60 predicted ABC Transporters in worms
classified MRP-4 as an ABCC Transporter. This subfamily includes the human
multidrug resistance proteins MRP1 to MRP-6; MRP-4 is most similar in sequence
to human MRP1 and MRP3 (Sheps et al.,
2004). Sequence analysis of cDNA clones identified an SL1 splice
leader sequence 43 nucleotides before the predicted start codon
(Krause and Hirsh, 1987)
(Fig. 1D). MRP-4 is predicted
to be 1573 amino acids long and to contain two ATP-binding cassettes. The
cd8 mutation results in an early stop codon right before the first
intron of the gene and is therefore predicted to be a null allele
(Fig. 1D).
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). We
also focused on developing intestinal cells at these embryonic stages because
we can unambiguously identify intestinal (and pharyngeal) cells in
cup-5(zu223) embryos based on position and morphology in the embryos.
We see clear suppression of the lysosomal defects and we see clear MRP-4
expression only in this tissue (see below).
We had previously shown that the degradation of at least two membrane
proteins, the yolk receptor RME-2 and the signaling protein LIN-12 is delayed
in cup-5(zu223) embryos (Fig.
2) (Schaheen et al.,
2006). By contrast, mrp-4(cd8); cup-5(zu223) embryos show
normal degradation of RME-2 and of LIN-12
(Fig. 2).
We also checked the lysosomal degradation of a soluble protein. Yolk, and
its associated proteins, is first endocytosed by oocytes and degraded by
embryonic cells during development. During embryonic morphogenesis, yolk is
secreted by all cells into the perivitelline space and is subsequently
endocytosed by developing intestinal cells where it is stored in lysosomes
(Bossinger and Schierenberg,
1996). At this stage in wild-type embryos, we can visualize the
localization of yolk using a fusion of the yolk protein YP170 to GFP
(Grant and Hirsh, 1999): all
of the YP170::GFP compartments stain with LysoTracker Red
(Fig. 3A)
(Schaheen et al., 2006). We
had previously shown that following this step, endocytosed YP170 accumulates
in LysoTracker Red-staining expanded `lysosomes' in cup-5(zu223)
embryos (Fig. 3A)
(Schaheen et al., 2006). By
contrast, 100% of the embryonic intestinal cells in both mrp-4(cd8)
and mrp-4(cd8); cup-5(zu223) had normal-sized YP170-containing and
LysoTracker Red-staining lysosomes (Fig.
3A,B). The average surface areas of YP170::GFP/LysoTracker
Red-staining compartments in confocal sections were 1.03±0.37
µm2 (wild type), 1.03±0.23 µm2
(mrp-4), 4.21±2.45 µm2 (cup-5) and
1.26±0.5 µm2 (mrp-4; cup-5). The sizes of these
compartments are essentially the same between wild type and mrp-4
(P 0.97), and between mrp-4 and mrp-4; cup-5
(P 0.16), but significantly different between wild type and
cup-5 cells (P 0.0003). The borders of the
YP170::GFP-staining compartments in all strains remains the same irrespective
of exposure time, indicating that it is not an artifact caused by the
saturation of the signal.
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mrp-4(cd8) rescues the developmental defects of cup-5(zu223)
If lysosomal function is restored, then we expected that developing
intestinal cells of mrp-4(cd8); cup-5(zu223) embryos would no longer
show evidence of starvation (Schaheen et
al., 2006). Indeed, in all of the mrp-4(cd8);
cup-5(zu223) embryos, intestinal cells no longer activate autophagy to
the same extent as cup-5(zu223) embryos, as determined by the
fluorescence intensity using a GFP::LGG-1 (MAP-LC3 ortholog) reporter
(Melendez et al., 2003;
Schaheen et al., 2006)
(Fig. 3C, large arrows). This
rescue is specific to intestinal cells, as we still see increased GFP::LGG-1
staining in other cells and is consistent with the intestine-specific
expression of MRP-4 (Fig. 3C,
see below). Consistent with this result, mrp-4(cd8); cup-5(zu223)
embryos, like cup-5(zu223) single mutants, still show increased
apoptosis relative to wild type (Fig.
3D) (Gonzalez-Polo et al.,
2005; Lum et al.,
2005; Takacs-Vellai et al.,
2005). We do not know whether the exacerbation of the lysosomal
defect in tissues other than intestinal cells of cup-5(zu223) embryos
is due to the accumulation of another ABC transporter or whether it is due to
distinct mechanisms.
Providing mrp-4(cd8); cup-(zu223) worms with methylpyruvate, a
membrane-permeable TCA substrate, slightly enhanced the number of hatched eggs
(97±3%; P 0.07) but had no effect on the viability of hatched
eggs (70±18%; P 0.8) or the sterility of the adults
(21±12%; P 0.27) (Fig.
1A-C). This is in agreement with previous results that showed that
methylpyruvate suppresses the autophagy and apoptosis defects but only
partially rescues embryonic and does not rescue larval lethality of
cup-5(zu223) worms by suppressing ectopic apoptosis
(Fig. 1A,B)
(Schaheen et al., 2006). The
major cause of lethality of cup-5(zu223) embryos is developmental
defects, including premature expression of the pmyo-2::GFP pharyngeal
reporter and abnormalities in intestinal tissue architecture, based on the
abnormal localization of the junctional intermediate filament protein IBF-2,
in 100% of cup-5(zu223) embryos: these are not rescued by
methylpyruvate (Schaheen et al.,
2006). In contrast to methylpyruvate, none of the mrp-4(cd8);
cup-5(zu223) show either one of these developmental defects
(Fig. 2,
Fig. 3C, small arrows). Thus,
restoration of lysosomal function of cup-5(zu223) embryos is
associated with suppression of developmental defects and with embryonic and
larval survival.
MRP-4 is required for the transport of lipophilic compounds that accumulate in the large vacuoles of cup-5(zu223) embryos
Cells from individuals with mucolipidosis Type IV accumulate lipids in
large vacuoles (Bargal and Bach,
1997; Chen et al.,
1998; Jansen et al.,
2001). We therefore asked whether cup-5(zu223) embryonic
intestinal cells also accumulate lipids in large vacuoles and whether this was
linked to the presence of MRP-4. Conjugates of lipophilic compounds with
glutathione are preferred physiological substrates for human MRP1, MRP2 and
MRP6 (Hirohashi et al., 1999;
Jedlitschky and Keppler, 2002;
Konig et al., 1999;
Leier et al., 1996;
Leier et al., 1994;
Zeng et al., 2000). We
therefore stained embryos with the lipophilic membrane-permeable dye Nile Red
to determine whether there is an increase in lipid stores in embryos. Nile Red
diffuses across membranes and fluoresces when it complexes with lipids
(Bossinger and Schierenberg,
1996; Greenspan et al.,
1985). We again used quantitative fluorescence measurements
instead of biochemical assays because the defect in cup-5(zu223)
embryos is confined to specific stages of embryonic development
(Schaheen et al., 2006).
In wild-type developing intestinal cells, Nile Red stains intracellular compartments that only occasionally overlap with the YP170::GFP-stained compartments. This indicates that Nile Red-staining lipids are mostly absent from the lysosomes of developing intestinal cells in wild-type embryos (Fig. 3E,F). The average integrated density (ID), which represents intensity per unit area of Nile Red-stained compartments is 0.03±0.017. mrp-4(cd8) results in a noticeable decrease in Nile Red staining (ID=0.004±0.0014; P 0.004, relative to wild type) and which, as in wild type, only occasionally overlaps with YP170::GFP (Fig. 3E,F). This indicates that MRP-4 probably transports the Nile Red-staining lipophilic substrates into endosomes/lysosomes.
cup-5(zu223) results in a substantial increase in Nile Red staining (ID=0.091±0.024; P 0.006, relative to wild type) (Fig. 3E,F). All of the enlarged compartments that contain YP170::GFP also stain for Nile Red. However, some Nile Red-stained compartments do not contain YP170::GFP (Fig. 3E). This indicates that the maturing and terminal endocytic compartments in cup-5(zu223) embryonic cells accumulate Nile Red-staining lipids. This increase in Nile Red staining is mostly due to MRP-4 activity, as opposed to the lack of CUP-5 function, as it is largely absent in mrp-4(cd8); cup-5(zu223) embryos (ID=0.0032±0.0019; P 0.004, relative to wild type; P 0.41, relative to mrp-4) (Fig. 3E,F).
MRP-4 is expressed in developing intestinal cells and accumulates in cup-5(zu223) embryos
To determine the expression pattern of MRP-4, we fused 3 kb of
mrp-4 promoter that includes sequences up to the next gene to GFP.
The introns in mrp-4 are small and unlikely to include important
regulatory elements. This transcriptional reporter showed that mrp-4
is expressed in developing intestinal cells at all stages of development,
which is consistent with earlier reports
(Zhao et al., 2004)
(Fig. 4A). This suggests that
intestinal defects contribute significantly to the death of embryos lacking
CUP-5. This is also consistent with the high incidence of sterility in rescued
mrp-4(cd8); cup-5(zu223) embryos and with the lack of rescue of the
lysosomal defect in cup-5(ar465) and cup-5(zu223)
coelomocytes by mrp-4(cd8), although we do not know yet whether these
defects are due to elevated levels of another ABC transporter
(Fares and Greenwald, 2001b;
Treusch et al., 2004;
Xue and Horvitz, 1997) (H.F.,
unpublished).
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MRP-4::GFP localizes to discrete compartments (integrated density of the GFP staining is 0.021±0.01) that partially co-localize with LysoTracker Red in developing intestinal cells. This indicates that at least some MRP-4::GFP localizes to late endocytic compartments. Some compartments stain with MRP-4::GFP but not with LysoTracker Red and may represent other endocytic compartments or cellular organelles. Conversely, some compartments stain with LysoTracker Red but not with MRP-4::GFP: these may constitute lysosomes that are devoid of MRP-4, either because MRP-4 is normally degraded in these compartments or because MRP-4 does not normally traffic to lysosomes but is transported to other organelles (Fig. 4C).
In cup-5(zu223) developing intestinal cells, MRP-4::GFP puncta are dramatically increased in size and intensity (integrated density of the GFP staining is 0.097±0.05, P 0.009, relative to wild type), indicating a defect in the degradation of MRP-4 (Fig. 4C). In this background, all of the LysoTracker Red-staining compartments contained MRP-4::GFP, indicating that this protein accumulates in the terminal cup-5(zu223) compartments. Some of the MRP-4::GFP-labeled enlarged compartments did not stain for LysoTracker Red in cup-5(zu223) embryos and may represent `maturing' endosomes/lysosomes intermediates (Fig. 4C). MRP-4::GFP staining almost completely overlaps with Nile Red both in wild-type and in cup-5(zu223) cells, which is consistent with the intensity of the Nile Red staining in endosomes/lysosomes being dependent on the presence of MRP-4 (Fig. 3E, Fig. 4D).
cup-5(zu223) lethality is not due to enlarged vacuoles/lysosomes
We thought it possible, although unlikely, that the loss of MRP-4 may
rescue the lethality and lysosomal function of cup-5(zu223) embryos
by reducing the sizes of lysosomes. We tested this indirectly by testing
whether mutations in upstream regulators of endocytosis rescue the lethality
of cup-5(zu223) embryos. Loss-of-function mutations in RME-1
(EH-domain protein required in recycling endosomes), RME-6 (RAB-5 GDP/GTP
exchange factor required at the plasma membrane and early endosomes) or CUP-10
(myotubularin required at the plasma membrane and endosomes) significantly
reduce lysosome sizes of cup-5(zu223) embryos
(Dang et al., 2003;
Grant et al., 2001;
Sato et al., 2005;
Xue et al., 2003) (see Fig. S1
in the supplementary material). None of these mutations rescue the
accumulation of Nile Red-staining compounds in compartments of developing
intestinal cells in cup-5(zu223) embryos (see Fig. S2 in the
supplementary material). This is in effect the reverse phenotypes of what we
see for mrp-4(cd8). However, none of these mutations rescued the
lethality of cup-5(zu223) embryos, indicating that reducing the rate
of uptake, which leads to a reduction in the sizes of lysosomes, is not
sufficient to alleviate the lethality of cup-5(zu223) (see Fig. S3 in
the supplementary material).
Rescue of cup-5(zu223) is specific to reducing levels of MRP-4
We used RNAi to test whether reducing the levels of other ABC transporters
in worms rescues the cup-5(zu223) embryonic lethality. RNAi of
mrp-4 results in 65.5±2% of the cup-5(zu223) embryos
hatching, indicating that this is a feasible approach to test the specificity
of the suppression by MRP-4. Of the 59 remaining predicted worm ABC
transporters, only RNAi of pgp-2 gave any suppression of embryonic
lethality. This was surprising as PGP-2 is not even expressed in the intestine
(Nunes et al., 2005). Indeed,
the null mutation pgp-2(gk114) did not produce any rescue of
cup-5(zu223) (Nunes et al.,
2005). The rescue by pgp-2 RNAi is probably due to
sequences in the pgp-2 double-stranded RNA that are complementary to
sequences in other worm ABC transporters, including MRP-4. These results
indicate that suppression of cup-5(zu223) lethality is not a general
feature of reducing ABC transporter levels, but is specific to MRP-4. Of the
27 worm ABC transporters that are expressed in intestinal cells, PGP-1 and
PGP-3 localize to the apical membrane, whereas nothing is known about the
subcellular distribution of the rest
(Broeks et al., 1995;
Zhao et al., 2004).
Loss of MRP-4 does not rescue lethality due to reduced endocytic uptake or to activation of apoptosis
The reversal of the cup-5(zu223) lysosomal degradation defect by
the loss of MRP-4 suggests that mrp-4(cd8) would not rescue embryonic
lethality because of non-lysosomal defects. Indeed, mrp-4(cd8) does
not rescue a loss-of-function mutation in ced-9, which results in the
activation of apoptosis in most embryonic cells
(Fig. 5A). Furthermore,
mrp-4(cd8) does not rescue the lethality caused by a null mutation in
the yolk receptor RME-2, which severely reduces the availability of yolk in
embryonic cells without affecting lysosomal function
(Fig. 5A).
Loss of MRP-4 partially rescues lethality due to loss of cathepsin L
We finally asked whether loss of MRP-4 could rescue lysosomal defects that
are due to the loss of other proteins than CUP-5. A null mutation in CPL-1,
the cathepsin L protease, results in progressively enlarged lysosomes and
embryonic lethality (Britton and Murray,
2004). Loss of MRP-4 does not rescue this lethality
(Fig. 5A). The caveat in this
experiment is that embryos lacking CPL-1 die at a much earlier stage than the
onset of MRP-4 expression in intestinal cells
(Britton and Murray, 2004). It
had previously been shown that reducing the levels of RME-2 (the yolk
receptor) allows some cpl-1 mutant embryos to develop to later stages
(Britton and Murray, 2004). We
thought that these embryos may arrest at the later stages because of the
accumulation of MRP-4 in the absence of CPL-1. We therefore reduced the levels
of RME-2 by RNAi in cpl-1 and in mrp-4; cpl-1 mutants. We
consistently saw a small fraction of embryos that hatched and that developed
to adults after RNAi of mrp-4; cpl-1 but not of cpl-1
mutants (Fig. 5B). This
suggests that increased MRP-4 activity, either resulting from the absence of
the transmembrane channel CUP-5 or from the loss of the lumenal enzyme CPL-1,
exacerbates their defects.
|
DISCUSSION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Salameh,
2006). The suppression of the premature pharyngeal expression is
probably non-cell-autonomous, and could be due either to defective connections
between intestinal and pharyngeal tissues and/or to signaling abnormalities
between the two tissues in the absence of CUP-5.
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Several mammalian ABC transporters localize to late endosomes/lysosomes,
including the ABCB transporter (MDR/TAP) member 6, ABCA1, ABCA2, ABCA3, ABCA5,
ABCB9 and the ABCC transporter MRP2
(Bagshaw et al., 2005;
Chen et al., 2005;
Cheong et al., 2006;
Kubo et al., 2005;
Neufeld et al., 2004;
Zhang et al., 2000;
Zhou et al., 2001). ABCA1
traffics between late endosomes and the cell surface and is required for the
efflux from the cell of cholesterol in late endosomes
(Chen et al., 2005;
Neufeld et al., 2004).
ABCA5-knockout mice accumulate lysosomes and autophagosomes in their
cardiomyocytes, indicating a requirement of these transporters in late
endocytic trafficking (Kubo et al.,
2005). Finally, and consistent with the proposed function of MRP-4
in the transport of lipophilic substances into compartments, ABCA3 is required
for late endosomal/lysosomal loading of phosphatidylcholine and for the
maturation of late endosomes (Cheong et
al., 2006).
Distinct pathways may be the source of this lysosome `overload', and hence
the exacerbation of lysosomal defects resulting from a primary lesion, in
different cell types. In developing intestinal cells of worm embryos, MRP-4
transports compounds into endosomes/lysosomes. In other cells, lysosome
`overloading' may be accomplished by endocytosis of various substrates. For
example, embryos laid by worms lacking CPL-1, the cathepsin L protease, are
unable to degrade both the yolk protein YP170 and yolk itself
(Britton and Murray, 2004).
Furthermore, decreasing the lipid load in lysosomes of this mutant, by
reducing the amount of yolk taken up by oocytes, reduces the size of the
expanded lysosomes and allows some of these embryos to develop to later stages
(Britton and Murray,
2004).
Most lysosomal disorders show similar cell biological abnormalities:
lysosomal enlargement and the accumulation of multiple substrates
(Vellodi, 2005). We propose
that a common mechanism for the exacerbation of the symptoms is that a primary
lesion results in the progressive accumulation of substrates in lysosomes that
ultimately lead to inhibition of other lysosomal enzymes. In some cell types,
a reduction in the rate of degradation of endosomal/lysosomal ABC transporters
may be the primary reason for the accumulation of these substrates.
Therapies for lysosomal storage diseases based on supplying cells with the
enzymes they are deficient for do not result in complete recovery with results
that vary considerably based on the age of onset and the rapidity of
progression (Desnick, 2004;
Jeyakumar et al., 2005;
Vellodi, 2005). We propose
that in these more severe cases, the lysosomes may have already accumulated
substances that inactivate lysosomal enzymes. New enzymes that are delivered
to these aberrant lysosomes may not be fully functional. Inactivating
endosomal/lysosomal ABC transporters, in conjunction with established
therapies, may prove more effective for restoring lysosomal function of some
cells.
There is considerable research aimed at inactivating MDR and MRP
transporters because of their ability to confer resistance to cytotoxic and
antiviral drugs (Glavinas et al.,
2004; Haimeur et al.,
2004). Recent studies using diterpenic lactones isolated from
Euphorbia, or shRNAi constructs, have shown a reversal of the
multidrug resistance activity of the ABCB transporter MDR1 in mouse cells
(Ferreira et al., 2005;
Pichler et al., 2005).
Reducing the activity of lysosomal ABC transporters in various cells may be a
viable treatment strategy for the treatment of mucolipidosis type IV, and
perhaps of other lysosomal disorders.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/133/19/3939/DC1
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ACKNOWLEDGMENTS |
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0.01
µm2. (C) Confocal micrographs of `1.5-fold' stage embryos
laid by the indicated hermaphrodites carrying the GFP::LGG-1 transgene. Long
arrows indicate staining of intestinal cells. This transgene also carries a
marker that expresses GFP in pharyngeal cells (short arrows). All images were
taken with the same exposure and at the same magnification. (D)
Confocal micrographs of wild-type, mrp-4(cd8), cup-5(zu223)
or mrp-4(cd8); cup-5(zu223) `comma' stage embryos stained using the
TUNEL assay to detect DNA-strand breaks. The average number of TUNEL-staining
bodies in each strain is indicated at the top of each panel. (E)
Confocal micrographs of `1.5 fold' stage embryos laid by hermaphrodites
carrying the YP170::GFP transgene (green in merge) and stained for Nile Red
(red in merge). All images of the same marker were taken with the same
exposure and at the same magnification. Arrows indicate staining of intestinal
cells. (F) Quantitation of the intensity per surface area of the Nile
Red granules in intestinal cells shown in E. Scale bars: 10 µm in all
images. The alleles in all panels are mrp-4(cd8) and
cup-5(zu223).
