REPORTS
`
`30 hours after infection of mosquitoes lacking
`LRIM1, APL1, or TEP1. As reported previously
`(16), three distinct classes of parasites were ob-
`served in the midguts of control mosquitoes: live
`(GFP positive), dead (TEP1 positive), and dying
`(GFP and TEP1 positive). In contrast, TEP1-positive
`parasites were never detected in LRIM1 or APL1
`KD mosquitoes (Fig. 3C), despite the presence
`of TEP1-F in the hemolymph. Lack of TEP1 par-
`asite staining was as complete as in TEP1 KD
`mosquitoes, which entirely lack TEP1 in the hemo-
`lymph. These data demonstrate that the LRIM1/
`APL1C complex is necessary for TEP1-mediated
`parasite killing during midgut invasion and in-
`dicate that TEP1 binds parasites only after it is
`processed.
`The APL1 locus has been implicated in
`mosquito resistance to the human malaria para-
`site, Plasmodium falciparum (6), and TEP1 has
`been shown to act against P. falciparum in labo-
`ratory infections (7). Mosquito defense against
`Plasmodium is likely to be influenced by vector/
`parasite coevolution and adaptation; thus, the ob-
`servation that LRIM1 did not affect P. falciparum
`in experimental field infections (17) may suggest
`that parasites have evolved to evade this pathway.
`Proteins such as the fibrinogen-related FBN9 (18)
`or other LRR proteins may provide alternative
`mechanisms for TEP1-mediated parasite killing.
`Bioinformatic searches for proteins related to
`LRIM1 and APL1C using their shared structural
`features (signal peptide, LRRs, cysteine pattern,
`and coiled-coils) (see supporting online material)
`detected more than 20 LRIM-like genes in each
`of the available mosquito genomes—A. gambiae,
`Aedes aegypti, and Culex quinquefasciatus—but
`not in any other species (table S1). Several of these
`genes were previously implicated in A. gambiae
`immune responses (3, 6, 7, 9, 19). Phylogenetic
`analysis in conjunction with pairwise compar-
`isons, examination of orthologous genomic neigh-
`borhoods, and protein domain analysis revealed
`four distinct LRIM subfamilies (figs. S3 and S4).
`Thus, LRIM1 and APL1C are members of a
`family of putative recognition receptors, which
`appears to be unique and greatly expanded in
`mosquitoes. Nevertheless, structural integrity of
`both LRRs and coiled-coils rests with only a few
`key amino acids, allowing considerable sequence
`variation that may hinder identification of func-
`tional equivalents in other organisms.
`The versatile LRR motif mediates recognition
`of diverse pathogen-associated molecules in host
`innate defense in plants and animals (20). For
`example, the repertoire of variable lymphocyte
`receptor (VLR) antibodies in jawless vertebrates
`is generated by combinatorial assembly of
`LRR modules instead of immunoglobulin seg-
`ments as in jawed vertebrates (21). Similarly to
`LRIM1/APL1C, the VLR antibodies are secreted
`as disulfide-linked multimers (22).
`LRIMs form a family of mosquito LRR re-
`ceptors with putative roles in defense against hu-
`man and animal pathogens. LRIM1 and APL1C
`exist as a complex that mediates immunity
`
`against malaria parasites through activation of
`mosquito complement. The multimeric nature of
`the complex indicates the potential to bind mul-
`tiple targets similarly to mammalian multisubunit
`receptors that robustly activate complement, that
`is, immunoglobulin M, lectin, and C1q. Bound
`LRIM1/APL1C complex may then undergo con-
`formational changes inducing the recruitment of
`additional cascade components, such as a TEP1-
`activating protease. In-depth study of these in-
`teractions will provide insights into complement
`activation in mosquitoes and tools toward block-
`ing disease transmission.
`References and Notes
`1. R. W. Snow, C. A. Guerra, A. M. Noor, H. Y. Myint, S. I. Hay,
`Nature 434, 214 (2005).
`2. S. A. Blandin, E. Marois, E. A. Levashina, Cell Host
`Microbe 3, 364 (2008).
`3. M. A. Osta, G. K. Christophides, F. C. Kafatos, Science
`303, 2030 (2004).
`4. E. Warr, L. Lambrechts, J. C. Koella, C. Bourgouin,
`G. Dimopoulos, Insect Biochem. Mol. Biol. 36, 769
`(2006).
`5. L. F. Moita et al., Immunity 23, 65 (2005).
`6. M. M. Riehle et al., Science 312, 577 (2006).
`7. Y. Dong et al., PLoS Pathog. 2, e52 (2006).
`8. T. Habtewold, M. Povelones, A. M. Blagborough,
`G. K. Christophides, PLoS Pathog. 4, e1000070 (2008).
`9. M. M. Riehle et al., PLoS One 3, e3672 (2008).
`10. F. H. Collins et al., Science 234, 607 (1986).
`11. G. Dimopoulos, A. Richman, H. M. Muller, F. C. Kafatos,
`Proc. Natl. Acad. Sci. U.S.A. 94, 11508 (1997).
`12. H. M. Muller, G. Dimopoulos, C. Blass, F. C. Kafatos,
`J. Biol. Chem. 274, 11727 (1999).
`
`13. S. Blandin et al., Cell 116, 661 (2004).
`14. E. A. Levashina et al., Cell 104, 709 (2001).
`15. R. H. Baxter et al., Proc. Natl. Acad. Sci. U.S.A. 104,
`11615 (2007).
`16. C. Frolet, M. Thoma, S. Blandin, J. A. Hoffmann,
`E. A. Levashina, Immunity 25, 677 (2006).
`17. A. Cohuet et al., EMBO Rep. 7, 1285 (2006).
`18. Y. Dong, G. Dimopoulos, J. Biol. Chem. www.jbc.org/cgi/
`doi/10.1074/jbc.M807084200. Published online
`4 February 2009.
`19. R. Aguilar et al., Insect Biochem. Mol. Biol. 35, 709 (2005).
`20. T. Nurnberger, F. Brunner, B. Kemmerling, L. Piater,
`Immunol. Rev. 198, 249 (2004).
`21. Z. Pancer et al., Nature 430, 174 (2004).
`22. B. R. Herrin et al., Proc. Natl. Acad. Sci. U.S.A. 105,
`2040 (2008).
`23. The authors thank A. C. Koutsos for generating the LRIM1
`antibody and sharing it before publication and F. M. Ausubel
`for critically reviewing the manuscript. Fluorescence
`microscopy was performed at the Imperial College Facility for
`Imaging by Light Microscopy imaging facility. This work
`was supported by a Wellcome Trust Program grant
`(GR077229/Z/05/Z), an NIH Program Project
`(2PO1AI044220-06A1), and a Biotechnology and Biological
`Sciences Research Council grant (BB/E002641/1). R.M.W.
`was supported by a Wellcome Trust Ph.D. fellowship.
`Supporting Online Material
`www.sciencemag.org/cgi/content/full/1171400/DC1
`Materials and Methods
`Figs. S1 to S4
`Table S1
`References
`
`26 January 2009; accepted 24 February 2009
`Published online 5 March 2009;
`10.1126/science.1171400
`Include this information when citing this paper.
`
`Glioma-Derived Mutations in IDH1
`Dominantly Inhibit IDH1 Catalytic
`Activity and Induce HIF-1a
`Shimin Zhao,1,2 Yan Lin,1* Wei Xu,1,2* Wenqing Jiang,1,2* Zhengyu Zha,1 Pu Wang,1,2
`Wei Yu,1,2 Zhiqiang Li,4 Lingling Gong,5 Yingjie Peng,6 Jianping Ding,6 Qunying Lei,1,3
`Kun-Liang Guan,1,3,7† Yue Xiong1,2,8†
`Heterozygous mutations in the gene encoding isocitrate dehydrogenase-1 (IDH1) occur in certain
`human brain tumors, but their mechanistic role in tumor development is unknown. We have shown
`that tumor-derived IDH1 mutations impair the enzyme’s affinity for its substrate and dominantly
`inhibit wild-type IDH1 activity through the formation of catalytically inactive heterodimers.
`Forced expression of mutant IDH1 in cultured cells reduces formation of the enzyme product,
`a-ketoglutarate (a-KG), and increases the levels of hypoxia-inducible factor subunit HIF-1a, a
`transcription factor that facilitates tumor growth when oxygen is low and whose stability is
`regulated by a-KG. The rise in HIF-1a levels was reversible by an a-KG derivative. HIF-1a levels
`were higher in human gliomas harboring an IDH1 mutation than in tumors without a mutation.
`Thus, IDH1 appears to function as a tumor suppressor that, when mutationally inactivated,
`contributes to tumorigenesis in part through induction of the HIF-1 pathway.
`
`Gliomas are the most common type of
`
`human brain tumors and can be classified
`based on clinical and pathological criteria
`into four grades. The grade IV glioma, commonly
`known as glioblastoma multiforme (GBM), has
`one of the worst prognoses among all types of
`human tumors and can develop either de novo
`(primary GBM) or through progression from low-
`grade tumors (secondary GBM). Although patho-
`
`logically indistinguishable, primary and secondary
`GBM exhibit distinct patterns of cancer gene al-
`terations (1). A recent cancer genome sequencing
`project revealed that the gene encoding IDH1 is
`somatically mutated predominantly in secondary
`GBM (2). Three subsequent studies of targeted
`IDH1 gene sequencing confirmed this finding,
`together identifying IDH1 mutations in more than
`70% of secondary GBM or low-grade gliomas but
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`infrequently in primary GBM (about 5%) (3–5).
`Notably, all of the IDH1 mutations identified to
`date produce a single amino acid substitution at
`Arg132 (R132) and no obvious inactivating (frame-
`shift or protein-truncation) mutations were found.
`This observation, together with the fact that the
`tumors do not show loss-of-heterozygosity (LOH),
`has led to speculation that R132 mutations lead to
`oncogenic activation of the enzyme.
`IDH enzymes catalyze the oxidative decar-
`boxylation of isocitrate (ICT) to produce a-KG.
`The human genome has five IDH genes coding
`for three distinct IDH enzymes whose activities
`are dependent on either nicotinamide adenine di-
`nucleotide phosphate (NADP+-dependent IDH1
`and IDH2) or nicotinamide adenine dinucleo-
`tide (NAD+-dependent IDH3). Both IDH2 and
`IDH3 enzymes are localized in the mitochon-
`dria and participate in the citric acid (TCA) cycle
`for energy production, whereas IDH1 is localized
`in the cytoplasm and peroxisomes (6). The R132
`residue is conserved in all NADP+-dependent
`IDHs.
`To explore the functional impact of the tumor-
`associated mutations at R132, we performed mod-
`eling studies based on the previously reported
`human cytosolic IDH1 crystal structure (7).
`Among all residues involved in binding with
`ICT, the side chain of R132 uniquely forms three
`hydrogen bonds with both the a- and b-carboxyl
`groups of the substrate ICT, whereas other resi-
`dues involved in ICT binding form no more than
`two hydrogen bonds (Fig. 1A). Substitution of
`R132 with any one of the six amino acids ob-
`served in gliomas (His, Ser, Gly, Cys, Val, and
`Leu) would impair interactions of the enzyme with
`ICT both sterically and electrostatically. Repre-
`sentative modeling of H132, which corresponds to
`the most prevalent IDH1 mutation in human
`gliomas, is shown in Fig. 1A. We determined the
`in vitro enzymatic activities of three tumor-derived
`IDH1 mutants, R132H, R132C and R132S, ex-
`pressed in transformed human embryonic kidney
`(HEK) 293T cells and found that all three mutants
`have a greater than 80% reduction in activity
`as compared with the wild-type (WT) IDH1
`(Fig. 1B). Analysis of recombinant IDH1 mutant
`
`1Molecular and Cell Biology Laboratory, Institute of Biomedical
`Sciences, Fudan University, 130 Dong-An Road, Shanghai
`200032, China. 2School of Life Sciences, Fudan University, 220
`Han-Dan Road, Shanghai 200433, China. 3Department of
`Biological Chemistry, School of Medicine, Fudan University,
`130 Dong-An Road, Shanghai 200032, China. 4Department of
`Neurosurgery, Zhongnan Hospital, Wuhan University, Wuhan
`430071, China. 5Department of Pathology, Zhongnan
`Hospital, Wuhan University, Wuhan 430071, China. 6State
`Key Laboratory of Molecular Biology, Institute of Biochemistry
`and Cell Biology, Shanghai Institute for Biological Sciences,
`Chinese Academy of Sciences, 320 Yue-Yang Road, Shanghai
`200031, China. 7Department of Pharmacology and Moores
`Cancer Center, University of California San Diego, La Jolla, CA
`92093, USA. 8Department of Biochemistry and Biophysics,
`Lineberger Comprehensive Cancer Center, University of North
`Carolina at Chapel Hill, NC 27599, USA.
`*These authors contributed equally to this work.
`†To whom correspondence should be addressed. E-mail:
`kuguan@ucsd.edu (K.-L.G.); yxiong@email.unc.edu (Y.X.)
`
`proteins purified from Escherichia coli likewise
`displayed little activity in vitro compared with
`wild-type controls (Fig. 1B). Kinetic analyses of
`recombinant IDH1 proteins revealed that all three
`mutant IDH1s had a dramatically reduced affini-
`ty for ICT: The Michaelis constants (Kms) of
`IDH1R132C, IDH1R132S, and IDH1R132H for ICT
`were increased by factors of 60, 70, and 94, re-
`spectively (Fig. 1C). In contrast, the Km for
`NADP+ and the maximum velocity (Vmax) were
`not appreciably altered (Fig. 1C). Similarly,
`mutation of an arginine residue in pig mitochon-
`drial IDH2 equivalent to R132 in human IDH1
`caused a dramatic increase in Km for isocitrate
`(by a factor of 165), with minimal effect on Vmax
`(8). Because the normal cellular concentration
`of ICT is 20 to 30 mM (9), which is lower than
`the Km of the IDH1 mutants, the mutant en-
`zymes are likely to have limited activity under
`physiological conditions. Together, these struc-
`tural and biochemical analyses indicate that
`the tumor-associated IDH1 mutations inactivate
`the enzyme.
`Given that IDH1 normally functions as a
`homodimer, we hypothesized that the mutant
`IDH1 molecules in tumor cells form heterodimers
`with wild-type molecules and, in so doing, domi-
`nantly inhibit the activity of wild-type IDH1. To
`test this hypothesis, we coexpressed His-tagged
`wild-type and FLAG-tagged R132H mutant
`IDH1 in E. coli and isolated the heterodimer by
`
`sequential affinity purification using first nickel
`resin and then FLAG beads. Formation of either
`WT:WT homodimer or WT:R132H heterodimer
`was confirmed by gel filtration (fig. S1). As ex-
`pected,
`the WT:WT homodimers were fully
`active and the R132H:R132H homodimers were
`nearly completely inactive (Fig. 2A). Notably, the
`WT:R132H heterodimer exhibited only 4% of the
`activity shown by the wild-type enzyme when
`assayed with limited ICT concentration (Fig. 2A).
`Normally, IDH1 can adopt at least three distinct
`conformations during catalysis: a quasi-open con-
`formation when it is in a complex with NADP+, a
`quasi-closed conformation when it is in a com-
`plex with ICT, and a closed conformation when it
`is in a complex with both NADP+ and ICT (fig.
`S2A) (7). The two IDH1 subunits act in a co-
`operative manner and undergo conformational
`changes in a concerted way. Our modeling study
`suggests that the impairment in enzyme binding
`with ICT conferred by the R132 mutation in one
`subunit might also impair the binding of ICT to
`the second wild-type subunit. As a result, both
`subunits would be locked in an unliganded or
`quasi-open (NADP+-bound) conformation,
`thereby inhibiting catalytic activity (Fig. 2B for
`the close-up of the catalytic active site) (fig. S2,
`B and C). Consistent with this model, we found
`that the wild-type IDH1 exhibits a sigmoidal
`curve of cooperative binding to ICT, whereas
`the WT:R132H heterodimer displayed a hy-
`
`Fig. 1. Tumor-derived IDH1 mutants have reduced catalytic activity because of impaired isocitrate
`binding. (A) Structural modeling predicts that mutation of R132 in IDH1 would weaken hydrogen bonding
`of the enzyme to ICT. Shown is a view of the catalytic active site of human IDH1 bound with NADP+
`(omitted for clarity), ICT (green), and Ca2+. The residues interacting with ICT from the adjacent subunit are
`labeled with an apostrophe. Hydrogen-bonding interactions are indicated with dashed lines. Simulated
`H132 mutation (cyan) is superimposed on R132. (B) Tumor-derived IDH1 mutants have reduced catalytic
`activity in vitro. Left, FLAG-tagged wild-type and mutant IDH1 were expressed in HEK293T cells, purified
`by immunoprecipitation and eluted by FLAG peptide; right, HIS-tagged wild-type and mutant IDH1 were
`expressed in E. coli and purified by nickel resin. Specific IDH1 activities for all proteins were measured in
`the presence of NADP+ (10 mM) and ICT (30 mM), with the presence of 2 mM Mn2+. Shown are mean
`values of triplicate experiments TSD. (C) Kinetic parameters of wild-type and mutant IDH1. Shown are
`mean values of duplicate experiments TSD.
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`perbolic curve with a much higher Km (Fig. 2C),
`indicating that the heterodimer not only loses
`affinity but also cooperativity toward ICT.
`We next investigated whether loss of IDH1
`activity would alter cellular levels of a-KG, the
`product of IDH1 catalysis. We used RNA in-
`terference to down-regulate endogenous IDH1
`and determined the a-KG levels in U-87MG
`
`human glioblastoma cells. Two independent
`short hairpin RNAs (shRNAs) decreased IDH1
`mRNA by more than 75% and reduced cellular
`a-KG levels by up to 50% (fig. S3). Expression
`of the IDH1R132H mutant at a level similar to the
`endogenous protein (fig. S4A) in the cytoplasm
`of U-87MG cells caused a dose-dependent re-
`duction of a-KG levels (Fig. 2D) (fig. S4, A and
`
`these data indicate that tumor-
`B). Together,
`derived mutant IDH1 dominantly inhibits the
`wild-type IDH1 by forming a catalytically in-
`active heterodimer, resulting in a decrease of
`cellular a-KG.
`Because a-KG is required by prolylhydrox-
`ylases (PHD), enzymes that hydroxylate and
`promote the degradation of hypoxia-inducible
`
`REPORTS
`
`Fig. 2. The R132H mutation dominantly inhibits
`IDH1 activity and reduces cellular levels of a-KG.
`(A) The WT:R132H heterodimer of IDH1 has low
`specific activity. The specific activities of WT:WT,
`R132H:R132H, and WT:R132H dimers were mea-
`sured under conditions of NADP+ (10 mM), ICT (30 mM),
`and 2 mM MnCl2. Activities were normalized by pro-
`tein levels, and wild-type activity was arbitrarily set as
`100%. Shown are mean values of triplicate experi-
`ments TSD. (B) A close-up view showing the con-
`formational differences between the IDH1-NADP+
`and IDH1-NADP+-ICT complexes at the active site.
`The enzyme adopts a quasi-open conformation in the
`IDH1-NADP+ complex (cyan) and a closed conforma-
`tion in the IDH1-NADP+-ICT complex (yellow). The
`bound ICT and the side chains of several residues in
`IDH1 involved in ICT binding are shown. (C) The
`WT:R132H heterodimer loses cooperative binding
`to ICT. The activities of the WT:WT and WT:R132H
`enzymes were assayed with increasing concentra-
`tions of ICT in the presence of 100 mM NADP+ and
`2 mM Mn2+. Shown are mean values of duplicate
`assays TSD. The inset is an expanded view showing
`the IDH1 activities at lower isocitrate concentrations.
`(D) Cellular a-KG levels decrease with increasing
`IDH1R132H expression. The upper panel is a Western
`blot showing expression levels of the transfected
`IDH1R132H mutant in U-87MG cells. The a-KG level
`in cells transfected with empty vector was set as
`100%, and this value was used to calculate the
`relative a-KG level in cells transfected with differ-
`ent amounts of IDH1R132H mutant. Shown are mean
`values of triplicate assays TSD.
`
`Fig. 3. a-KG mediates the HIF-1a induction in cells with a decreased
`IDH1 activity. (A) IDH1 knockdown elevates HIF-1a levels in U-87MG
`glioblastoma cells. IDH1 and HIF-1a protein levels were determined by
`Western blotting from stable U-87MG cells transduced with empty
`retrovirus or retrovirus expressing different shRNAs silencing IDH1. (B)
`Ectopic expression of the IDH1R132H mutant elevates HIF-1a levels in U-
`87MG and HEK293T cells. The IDH1R132H mutant was overexpressed in
`U-87MG or HEK293T cells, and protein levels were detected by Western
`blot. CoCl2-treated cells (a mimetic of hypoxia) and cells overexpressing
`wild-type IDH1 were also included as controls. (C) A cell-permeable a-KG
`derivative blocks HIF-1a induction in cells expressing IDH1R132H. The U-
`87MG cells were transfected with IDH1R132H, and different concen-
`trations of octyl-a-KG ester were added to each transfected cell for 4
`hours. HIF-1a protein levels were assayed by Western blot.
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`statistically stronger HIF-1a signal than did 12
`tumors that did not harbor this mutation (Fig.
`4C). In IDH1 mutated tumors, 28.1 T 6.7% of
`the cells stained positive for HIF-1a, whereas
`in tumors with wild-type IDH1, only 15.8 T
`3.5% of the cells stained positive (P < 0.001)
`(Fig. 4C). IDH1-mutated gliomas also exhibited
`an increase in VEGF levels compared with gliomas
`without IDH1 mutation of similar type and grade
`(fig. S7B).
`In summary, we have shown that IDH1 is
`likely to function as a tumor suppressor gene
`rather than as an oncogene. The glioma-associated
`mutations dominantly inhibit the activity of wild-
`type IDH1 through heterodimer formation (fig. S8).
`IDH1 gene mutations in gliomas exhibit
`two
`unique features: the lack of LOH and the lack
`of apparent inactivating mutations such as frame-
`shift or truncations. Our findings help to explain
`both features, as dominant
`inhibition would
`eliminate the selection pressure to mutate or lose
`the remaining wild-type allele (LOH) and frame-
`shift or premature termination would likely gen-
`erate IDH1 fragments unable to dimerize with
`and inhibit the wild-type IDH1 protein.
`The link between IDH1 and HIF-1a high-
`lights an emerging theme in which mutationally
`altered metabolic enzymes are thought to con-
`tribute to tumor growth by stimulating the HIF-1a
`pathway and tumor angiogenesis. The genes en-
`coding two TCA enzymes, fumarate hydratase
`(FH) and succinate dehydrogenase (SDH), have
`been found to sustain loss-of-function mutations
`in certain human tumors, which likewise corre-
`late with an increase in HIF-1a levels (10, 11). In
`addition to affecting PHD, an alteration in a-KG
`might contribute to tumorigenesis by affecting
`other dioxygenases that use a-KG as a substrate.
`IDH1 also catalyzes the production of NADPH;
`thus, it is possible that a reduction in NADPH
`levels resulting from IDH1 mutation contributes
`to tumorigenesis through effects on cell metabo-
`lism and growth. Several dozen anticancer agents
`directly targeting HIF-1a are under development
`or being tested (12). Our finding that an a-KG
`derivative can reverse the induction of HIF-1a
`levels in cultured cells expressing mutant IDH1
`suggests that drugs mimicking a-KG may merit
`exploration as a therapy for gliomas that harbor
`an IDH1 mutation.
`
`References and Notes
`1. F. B. Furnari et al., Genes Dev. 21, 2683 (2007).
`2. D. W. Parsons et al., Science 321, 1807 (2008).
`3. J. Balss et al., Acta Neuropathol. 116, 597 (2008).
`4. F. E. Bleeker et al., Hum. Mutat. 30, 7 (2009).
`5. H. Yan et al., N. Engl. J. Med. 360, 765 (2009).
`6. B. S. Winkler, N. DeSantis, F. Solomon, Exp. Eye Res. 43,
`829 (1986).
`7. X. Xu et al., J. Biol. Chem. 279, 33946 (2004).
`8. S. Soundar, B. L. Danek, R. F. Colman, J. Biol. Chem.
`275, 5606 (2000).
`9. K. R. Albe, M. H. Butler, B. E. Wright, J. Theor. Biol. 143,
`163 (1990).
`10. E. D. MacKenzie et al., Mol. Cell. Biol. 27, 3282
`(2007).
`11. O. C. Ingebretsen, Biochim. Biophys. Acta 452, 302
`(1976).
`
`Fig. 4. IDH1 activity affects the levels of HIF-1a and HIF-1a target genes in gliomas and cultured cells.
`(A) Overexpression of the IDH1R132H mutant in U-87MG cells stimulates expression of HIF-1a target genes
`(Glut1, VEGF, and PGK1) as assayed by QPCR. Shown are mean values of triplicate assays TSD. (B)
`Inhibition of IDH1 by oxalomalate activates HIF-1a target genes. U-87MG cells were either untreated
`(control), treated with 5 mM oxalomalate, an IDH1 inhibitor, or treated with CoCl2, a hypoxia mimetic.
`HIF-1a target gene mRNAs were determined. Shown are mean values of triplicate assays TSD. (C)
`Immunohistochemistry of HIF-1a was carried out in 12 human gliomas with wild-type IDH1 and 8 gliomas
`of similar grade harboring a mutated IDH1 allele. Shown are side-by-side comparisons of four gliomas
`representing different types or grades. Scale bar, 40 mM. Five fields (~173 mm2 each) were randomly
`selected from each sample for quantification of HIF-1a-positive staining area. Statistical analysis was
`performed using seven IDH1 wild-type and seven IDH1-mutated gliomas.
`
`factor 1a (HIF-1a), we hypothesized that de-
`creased IDH1 activity might stabilize HIF-1a and
`increase its steady-state levels. We found that
`HIF-1a protein levels in U-87MG cells were
`elevated in response to shRNA-mediated knock-
`down of IDH1 (Fig. 3A). Conversely, overexpres-
`sion of wild-type IDH1 reduced HIF-1a protein
`levels in HeLa (fig. S5A) and U-87MG cells
`(Fig. 3B). Notably, overexpression of IDH1R132H
`mutant increased HIF-1a protein levels in U-
`87MG and HEK293T cells (Fig. 3B). These
`results suggest that IDH1 regulates HIF-1a levels
`by controlling the level of a-KG. We tested this
`hypothesis by treating cells with octyl-a-KG, a
`cell-permeable derivative of a-KG that upon
`entering the cells is converted into a-KG after
`hydrolysis of the ester group (10). We found that
`octyl-a-KG suppressed the HIF-1a induction
`caused by either IDH1 knockdown in HeLa cells
`(fig. S5B) or overexpression of IDH1R132H mu-
`tant in U-87MG cells (Fig. 3C). We therefore con-
`clude that a reduction in IDH1 activity produces a
`reduction in a-KG levels that in turn can lead to
`stabilization of HIF-1a.
`We next determined whether inhibition of
`IDH1 enzyme activity leads to up-regulated ex-
`pression of HIF-1a target genes. HIF-1a is a key
`
`component of HIF-1, a transcription factor that
`senses low cellular oxygen levels and that regu-
`lates the expression of genes implicated in glucose
`metabolism, angiogenesis, and other signaling
`pathways that are critical to tumor growth. Quan-
`titative real-time fluorescence polymerase chain
`reaction (QPCR) of mRNAs corresponding to
`three well-established HIF-1a target genes, glu-
`cose transporter 1 (Glut1), vascular endothelial
`growth factor (VEGF), and phosphoglycerate
`kinase (PGK1) showed that IDH1 knockdown
`induced the expression of these HIF-1a target
`genes (fig. S6A). Moreover, expression of the
`IDH1R132H mutant, but not wild-type IDH1,
`strongly induced HIF-1a target gene expression
`(Fig. 4A). Oxalomalate, a competitive inhibitor
`of IDH1 (11) (fig. S6B), also induced expression
`of these HIF-1a target genes (Fig. 4B).
`Finally, we determined whether IDH1 muta-
`tions correlate with elevated levels of HIF-1a in
`human gliomas. In a collection of 26 glioma
`samples, we identified 8 tumors that contained
`the R132H mutation in one allele of IDH1 (table
`S1) (fig. S7A). Using immunohistochemistry, we
`compared HIF-1a expression in gliomas with
`and without IDH1 mutations. We found that 8
`tumors harboring the R132H mutation showed a
`
`264
`
`10 APRIL 2009 VOL 324 SCIENCE www.sciencemag.org
`
`Rigel Exhibit 1012
`Page 4 of 5
`
`

`

`12. G. L. Semenza, Drug Discov. Today 12, 853 (2007).
`13. We thank members of the Fudan Molecular and Cell
`Biology Laboratory for valuable input; Y. Liu, X. Liu, and
`H. Zhu for assistance with histology; Z. Bao, L. Yang,
`Q. Shi, and G. Zhao for clinical samples; and S. Jackson
`for reading the manuscript. This work is supported by the
`985 program from the Chinese Ministry of Education,
`State Key Development Programs of China
`
`(2009CB918401, 2006CB806700), National 863
`Program of China (2006AA02A308), China NSF grants
`(30600112 and 30871255) and Shanghai Key Basic
`Research Projects (06JC14086, 07PJ14011, and
`08JC1400900), and NIH grants (to K.-L.G. and Y.X.).
`Y. Xiong, K.-L. Guan, and S. Zhao are applying for
`a patent related to the work on permeable
`alpha-ketogluterate.
`
`Supporting Online Material
`www.sciencemag.org/cgi/content/full/324/5924/261/DC1
`Materials and Methods
`Figs. S1 to S8
`Table S1
`
`15 January 2009; accepted 10 March 2009
`10.1126/science.1170944
`
`REPORTS
`
`Demonstration of Genetic Exchange
`During Cyclical Development of
`Leishmania in the Sand Fly Vector
`Natalia S. Akopyants,1* Nicola Kimblin,2* Nagila Secundino,2 Rachel Patrick,2 Nathan Peters,2
`Phillip Lawyer,2 Deborah E. Dobson,1 Stephen M. Beverley,1† David L. Sacks2†‡
`
`Genetic exchange has not been shown to be a mechanism underlying the extensive diversity of
`Leishmania parasites. We report here evidence that the invertebrate stages of Leishmania are
`capable of having a sexual cycle consistent with a meiotic process like that described for African
`trypanosomes. Hybrid progeny were generated that bore full genomic complements from both
`parents, but kinetoplast DNA maxicircles from one parent. Mating occurred only in the sand fly
`vector, and hybrids were transmitted to the mammalian host by sand fly bite. Genetic exchange
`likely contributes to phenotypic diversity in natural populations, and analysis of hybrid progeny
`will be useful for positional cloning of the genes controlling traits such as virulence, tissue
`tropism, and drug resistance.
`
`Parasitic protozoa of the genus Leishmania
`
`cause a spectrum of human diseases that
`pose serious public health challenges for
`prevention, diagnosis, and treatment. The diver-
`sity of Leishmania species, with more than 20
`currently recognized, is thought to have arisen by
`gradual accumulation of divergent mutations rather
`than by sexual recombination. Tibayrenc et al.
`(1) have reported strong linkage disequilibrium
`in several Leishmania species and proposed that
`these parasites are essentially clonal. This notion
`must be reconciled, however, with the accumu-
`lating examples of naturally occurring strains that
`share genotypic markers from two recognized
`species and thereby provide circumstantial evi-
`dence for sexual recombination (2–4). Genetic
`exchange has been documented for the other
`trypanosomatids that cause human disease.
`Hybrid genotypes were observed in tsetse flies
`during cotransmission of two strains of Trypano-
`soma brucei (5) and in mammalian cells after
`coinfection with two clones of Trypanosoma
`cruzi differing in drug-resistance markers (6).
`Using drug resistance markers, we provide evi-
`dence for genetic exchange in Leishmania
`major and discuss the implications of these
`findings to Leishmania biology and experimen-
`tal analysis.
`
`1Department of Molecular Microbiology, Washington Univer-
`sity School of Medicine, St. Louis, MO 63110, USA. 2Labora-
`tory of Parasitic Diseases, National Institute of Allergy and
`Infectious Diseases, NIH, Bethesda, MD 20892, USA.
`*†These authors contributed equally to this work.
`‡To whom correspondence should be addressed. E-mail:
`dsacks@nih.gov
`
`One parental clone, LV39c5(HYG), was de-
`rived from strain LV39 clone 5 (MHOM/SU/59/P)
`and was heterozygous for an allelic replacement of
`the LPG5A on chromosome 24 by a hygromycin
`B–resistance cassette (LPG5A/LPG5A::ΔHYG)
`(7). The second parental clone, FV1(SAT), was
`derived from NIH Friedlin clone V1 (MHOM/
`IL/80/FN) and bore a heterozygous nourseothricin–
`resistance (SAT) marker, integrated along with a
`linked firefly luciferase (LUC) reporter gene into
`one allele of the ~24 rRNA cistrons located on
`chromosome 27 (8) (+/SSU::SAT-LUC). These
`strains were chosen as they are phenotypically
`identical to their respective parental wild-type (WT)
`virulent L. major; whereas the markers were chosen
`because they are functionally independent (9). The
`target gene modifications were chosen because
`they caused no effect on normal growth in vitro
`or in mouse infections (10), and epistatic interac-
`tions were not anticipated between these alleles.
`Multiple attempts to generate hybrid para-
`sites resistant to both antibiotics during in vitro co-
`culture of the parental lines were unsuccessful (11).
`The parental clones were tested for their ability to
`generate parasites resistant to both drugs during
`coinfection in the sand fly. The growth of each
`parental line in Phlebotomus duboscqi, a natural
`vector of L. major, is shown in fig. S1. Promas-
`tigotes of each parent survived the initial period
`of blood-meal digestion and excretion (days
`1 to 6) and underwent metacyclogenesis at a
`comparable frequency (20 to 60%), although the
`FV1(SAT) parent established and maintained a
`higher intensity of infection by a factor of 3 to 4.
`The parental clones were tested for their ability
`
`to generate doubly drug–resistant parasites dur-
`ing coinfection in the sand fly. Flies were fed
`through a membrane on mouse blood contain-
`ing 3 and 1 × 106/ml of the LV39c5(HYG) and
`FV1(SAT) lines, respectively, each obtained from
`log-phase cultures and extensively washed to
`remove antibiotics. A total of 102 flies from four
`independent coinfection experiments were dis-
`sected 13 to 16 days postinfection; at this time,
`they harbored mature infections with an average
`of 39,400 T 14,700 promastigotes per midgut.
`Flies cannot be maintained under aseptic con-
`ditions, and more than half of the cultures es-
`tablished from the midgut parasites were lost to
`fungal contamination during the subsequent 1 to
`2 weeks of culture. In the remaining cultures, 12
`(26%) grew out promastigotes that were resistant
`to both drugs. Clonal lines were generated from
`nine of the doubly drug–resistant populations,
`and the genotypes and phenotypes of one or two
`clones from each culture were determined (sum-
`marized in Table 1).
`Polymerase chain reaction (PCR) tests with
`primers specific for the parental markers showed
`that all doubly drug–resistant clones tested con-
`tained both the HYG and SAT drug-resistance
`genes (Fig. 1A, Table 1, and table S3). Controls
`showed that the marker loci had not rearranged
`dur