C11. Phosphatases - Biology

There are three main families of phosphatases, the phospho-Tyr phosphatases (PTP), the phospho-Ser/Thr phosphatases, and those that cleave both. Regulatory subunits for Tyr phosphatases may contain a SH2 domain allowing binding of the binary complex to autophosphorylated membrane receptor Tyr kinases.

Important Ser/Thr phosphatases (PPs for Protein Phosphatases) include:

  • Protein phosphatase 1 (PP-1 or Ppp1) - This is the most abundant PPPs in humans. Different regulatory subunits target this to the liver glycogen particles (GL subunit), striated muscle glycogen and sacroplasmic reticulum (GM subunit) or smooth muscle fibers (M subunit). It is also present in the nucleus where it is presumably involved in regulation of transcription factors. It is also involved in RNA splicing and signaling at neural synapses.
  • Protein phosphatase 2A (PP-2A or Ppp2) - is a trimer with catalytic, regulatory, and a scaffolding (also regulatory) structural subunits. It is found mainly in the cytoplasm and is involved in a myriad of cellular process.
  • Protein phosphatase 2B (PP-2B or Ppp3) - also called calcineurin or Ca2+/Calmodulin dependent protein phosphatase - It consists of a catalytic subunit (calcineurin A) and a regulatory, calcium-binding subunit (calcinerin B). It is inhibited by the complex of the immunosuppressant cyclosporin and FK506 with immunophilins. PP2B regulates PKA and PKC

PP1, 2A and 2B share a great deal of amino acid homology, and based on this homology, belong to one family. PP2C belongs to another. PPs are often categories into three families including, phosphoprotein phosphatases (PPPs) and metal-dependent protein phosphatases (PPMs). There about 30 catalytic PP subunits (many fold fewer than Ser/Thr Kinases). They gain specificity by binding numerous modulatory regulatory subunits.

As with other proteins, the names given to the proteins when discovered often do not reflect an organization scheme that would name different members based on structural similarities. PP-1, 2A, and 2B are better named Ppp1, Ppp2, and Ppp3 which denotes member of the Protein PP (PPP) family. PP-2C would be named Ppm1 as the first member of the PPN family. All PPPs have three short sequence motifs that bind divalent cations.

Protein Tyr phosphatases (PTPs) consist of receptor-like (transmembrane) and intracellular Tyr phosphatases. They more resemble tyrosine kinases in their complexity than the Ser/Thr phosphatases. There are about 100 PTPs in the genome, a number similar to the number of protein tyrosine kinases. PTPs have an active site Cys in a CX5R-(S/T) motif with an active site Cys nucleophile and an Arg in the phosphate binding (P) loop. Important examples include:

  • PTP1B - dephosphorylates many cell surface receptors (insulin, EGF, PDGF) that have been phosphorylated on Tyr residues. Its main activity seems to dephosphorylate nascent receptors in the endoplasmic reticulum before they get to the final cell membrane destination.
  • Low molecular weight PTPase - These have roles in metabolism and differentiation of cells. They have a molecular weight of 18,000 and have an active site CX5R-(S/T) motif, where the C (Cys) is an active site nucleophile.

Figure: PTP Super Family

Web Links for Phosphatases

Nontransmembrane and Receptor-Like Protein Tyrosine Phosphatases

Web Resources on Phosphatases


  • Prof. Henry Jakubowski (College of St. Benedict/St. John's University)

Adenosine diphosphate

Adenosine diphosphate (ADP), also known as adenosine pyrophosphate (APP), is an important organic compound in metabolism and is essential to the flow of energy in living cells. ADP consists of three important structural components: a sugar backbone attached to adenine and two phosphate groups bonded to the 5 carbon atom of ribose. The diphosphate group of ADP is attached to the 5’ carbon of the sugar backbone, while the adenine attaches to the 1’ carbon. [1]

  • 58-64-0 Y
  • CHEBI:16761 Y
  • ChEMBL14830 Y
  • 5800 Y
  • DB03431 N
  • C00008 N
  • 61D2G4IYVH Y
InChI=1S/C10H15N5O10P2/c11-8-5-9(13-2-12-8)15(3-14-5)10-7(17)6(16)4(24-10)1-23-27(21,22)25-26(18,19)20/h2-4,6-7,10,16-17H,1H2,(H,21,22)(H2,11,12,13)(H2,18,19,20)/t4-,6-,7-,10-/m1/s1 Y Key: XTWYTFMLZFPYCI-KQYNXXCUSA-N Y

ADP can be interconverted to adenosine triphosphate (ATP) and adenosine monophosphate (AMP). ATP contains one more phosphate group than does ADP. AMP contains one fewer phosphate group. Energy transfer used by all living things is a result of dephosphorylation of ATP by enzymes known as ATPases. The cleavage of a phosphate group from ATP results in the coupling of energy to metabolic reactions and a by-product of ADP. [1] ATP is continually reformed from lower-energy species ADP and AMP. The biosynthesis of ATP is achieved throughout processes such as substrate-level phosphorylation, oxidative phosphorylation, and photophosphorylation, all of which facilitate the addition of a phosphate group to ADP.


In humans there are 102 putative DUB genes, which can be classified into two main classes: cysteine proteases and metalloproteases, consisting of 58 ubiquitin-specific proteases (USPs), 4 ubiquitin C-terminal hydrolases (UCHs), 5 Machado-Josephin domain proteases (MJDs), 14 ovarian tumour proteases (OTU), and 14 Jab1/Mov34/Mpr1 Pad1 N-terminal+ (MPN+) (JAMM) domain-containing genes. 11 of these proteins are predicted to be non-functional, leaving 79 functional enzymes. [6] In yeast, the USPs are known as ubiquitin-specific-processing proteases (UBPs).

Cysteine proteases Edit

There are six main superfamilies of cysteine protease DUBs: [7]

  • the ubiquitin-specific protease (USP/UBP) superfamily (USP1, USP2, USP3, USP4, USP5, USP6, USP7, USP8, USP9X, USP9Y, USP10, USP11, USP12, USP13, USP14, USP15, USP16, USP17, USP17L2, USP17L3, USP17L4, USP17L5, USP17L7, USP17L8, USP18, USP19, USP20, USP21, USP22, USP23, USP24, USP25, USP26, USP27X, USP28, USP29, USP30, USP31, USP32, USP33, USP34, USP35, USP36, USP37, USP38, USP39, USP40, USP41, USP42, USP43, USP44, USP45, USP46)
  • the ovarian tumour (OTU) superfamily (OTUB1, OTUB2)
  • and the Machado-Josephin domain (MJD) superfamily. (ATXN3, ATXN3L)
  • the ubiquitin C-terminal hydrolase (UCH) superfamily (BAP1, UCHL1, UCHL3, UCHL5)
  • the MINDY family of K48-specific deubiquitinases (MINDY1, MINDY2, MINDY3, MINDY4) [8]
  • the recently discovered ZUFSP family, at present solely represented by ZUP1 [9]

There is also a little known putative group of DUBs called the permutated papain fold peptidases of dsDNA viruses and eukaryote (PPPDEs) superfamily, which, if shown to be bona fide DUBs, would be the seventh in the cysteine protease class. [10]

Metalloproteases Edit

The Jab1/Mov34/Mpr1 Pad1 N-terminal+ (MPN+) (JAMM) domain superfamily proteins bind zinc and hence are metalloproteases. [7]

DUBs play several roles in the ubiquitin pathway. One of the best characterised functions of DUBs is the removal of monoubiqutin and polyubiquitin chains from proteins. These modifications are a post translational modification (addition to a protein after it has been made) where single ubiquitin proteins or chains of ubiquitin are added to lysines of a substrate protein. These ubiquitin modifications are added to proteins by the ubiquitination machinery ubiquitin-activating enzymes (E1s), ubiquitin-conjugating enzymes (E2s) and ubiquitin ligases (E3s). The end result is ubiquitin bound to lysine residues via an isopeptide bond. [11] Proteins are affected by these modifications in a number of ways: they regulate the degradation of proteins via the proteasome and lysosome coordinate the cellular localisation of proteins activate and inactivate proteins and modulate protein-protein interactions. [3] [4] [5] DUBs play the antagonistic role in this axis by removing these modifications, therefore reversing the fate of the proteins. [2] In addition, a less understood role of DUBs is the cleavage of ubiquitin-like proteins such as SUMO and NEDD8. Some DUBs may have the ability to cleave isopeptide bonds between these proteins and substrate proteins. [12]

They activate ubiquitin by the proteolysis (breaking down) of the inactive expressed forms of ubiquitin. Ubiquitin is encoded in mammals by 4 different genes: UBA52, RPS27A, UBB and UBC. A similar set of genes is found in other eukaryotes such as yeast. The UBA52 and RPS27A genes produce ubiquitin that is fused to ribosomal proteins and the UBB and UBC genes produce polyubiquitin (a chain of ubiquitin joined by their C- and N-termini). [13] [14] DUBs cleave the ubiquitin from these proteins, producing active single units of ubiquitin. [2]

DUBs also cleave single ubiquitin proteins that may have had their C-terminal tails accidentally bound to small cellular nucleophiles. [2] These ubiquitin-amides and ubiquitin-thioesters may be formed during standard ubiquitination reactions by the E1-E2-E3 cascade. Glutathione and polyamines are two nucleophiles that might attack the thiolester bond between ubiquitin and these enzymes. Ubiquitin C-terminal hydrolase is an example of the DUB that hydrolyses these bonds with broad specificity. [12] [15]

Free polyubiquitin chains are cleaved by DUBs to produce monoubiquitin. The chains may be produced by the E1-E2-E3 machinery in the cell free from any substrate protein. Another source of free polyubiquitin is the product of ubiquitin-substrate cleavage. If DUBs cleave the base of the polyubiquitin chain that is attached to a protein, the whole chain will become free and needs to be recycled by DUBs. [2]

DUBs often contain a catalytic domain surrounded by one or more accessory domains, some of which contribute to target recognition. These additional domains include domain present in ubiquitin-specific proteases (DUSP) domain ubiquitin-like (UBL) domain meprin and TRAF homology (MATH) domain zinc-finger ubiquitin-specific protease (ZnF-UBP) domain zinc-finger myeloid, nervy and DEAF1 (ZnF-MYND) domain ubiquitin-associated (UBA) domain CHORD-SGT1 (CS) domain microtubule-interacting and trafficking (MIT) domain rhodenase-like domain TBC/RABGAP domain and B-box domain. [6] [16]

Catalytic domain Edit

The catalytic domain of DUBs is what classifies them into particular groups USPs, OTUs, MJDs, UCHs and MPN+/JAMMs. The first 4 groups are cysteine proteases, whereas the latter is a zinc metalloprotease. The cysteine protease DUBs are papain-like and thus have a similar mechanism of action. They use either catalytic dyads or triads (either two or three amino acids) to catalyse the hydrolysis of the amide bonds between ubiquitin and the substrate. The active site residues that contribute to the catalytic activity of the cysteine protease DUBs are cysteine (dyad/triad), histidine (dyad/triad) and aspartate or asparagine (triad only). The histidine is polarised by the aspartate or asparagine in catalytic triads or by other ways in dyads. This polarised residue lowers the pKa of the cysteine, allowing it to perform a nucleophilic attack on the isopeptide bond between the ubiquitin C-terminus and the substrate lysine. Metalloproteases coordinate zinc ions with histidine, aspartate and serine residues, which activate water molecules and allows them to attack the isopeptide bond. [17] [18]

UBL Edit

Ubiquitin-like (UBL) domains have a similar structure (fold) to ubiquitin, except they lack the terminal glycine residues. 18 USPs are proposed to have UBL domains. Only 2 other DUBs have UBLs outside the USP group: OTU1 and VCPIP1. USP4, USP7, USP11, USP15, USP32, USP40 and USP47 have multiple UBL domains. Sometimes the UBL domains are in tandem, such as in USP7 where 5 tandem C-terminal UBL domains are present. USP4, USP6, USP11, USP15, USP19, USP31, USP32 and USP43 have UBL domains inserted into the catalytic domain. The functions of UBL domains are different between USPs, but commonly they regulate USP catalytic activity. They can coordinate localisation at the proteasome (USP14) negatively regulate USPs by competing for the catalytic site of the USP (USP4), and induce conformational changes to increase catalytic activity (USP7). [16] [19] [20] Like other UBL domains, the structure of USP UBL domains show a β-grasp fold. [21] [22]


Single or multiple tandem DUSP domains of approximately 120 residues are found in six USPs. The function of the DUSP domain is currently unknown but it may play a role in protein-protein interaction, in particular to DUBs substrate recognition. This is predicted because of the hydrophobic cleft present in the DUSP domain of USP15 and that some protein interactions with DUSP containing USPs do not occur without these domains. The DUSP domain displays a novel tripod-like fold comprising three helices and an anti-parallel beta-sheet made of three strands. This fold resembles the legs (helices) and seat (beta-sheet) of the tripod. Within most DUSP domains in USPs there is a conserved sequence of amino acids known as the PGPI motif. This is a sequence of four amino acids proline, glycine, proline and isoleucine, which packs against the three-helix bundle and is highly ordered. [6] [23]

The full extent of the role of DUBs in diseases remains to be elucidated. Their involvement in disease is predicted due to known roles in physiological processes that are involved in disease states including cancer and neurological disorders. [24]

The enzyme USP28 is over-expressed in different types of cancer such as colon or lung. In addition, USP28 deubiquitinates and stabilizes important oncogenes such as c-Myc, Notch1, c-jun or ΔNp63. [25] [26] [27] In squamous tumors, USP28 regulates the resistance to chemotherapy regulating DNA repair via ΔNp63-Fanconia anemia pathway axis. [28]

The deubiquitinating enzymes UCH-L3 and YUH1 are able to hydrolyse mutant ubiquitin UBB+1 despite of the fact that the glycine at position 76 is mutated. [29]

UCH-L1 levels are high in various types of malignancies (cancer). [30]

DUBs play an active role in modulating the cell cycle. Ubiquitin-specific-processing protease (USP) is a family of deubiquitinating enzymes that play a crucial role in cell cycle regulation. [31] Two such enzymes include USP17 and USP44. USP17 regulates pathways responsible for progressing cells through the cell cycle. [32] Its targets include regulators of Ras, CDK2, and Cyclin A. [33] USP44 plays an important role in anaphase initiation. [34] New research into the mitotic checkpoint has revealed a novel role for USP44 in regulating cell cycle progression. [34]

USP Regulation of Ras Edit

The ERK Pathway allows for the transduction of external mitogenic signals into intracellular signals promoting cellular proliferation. One of the key regulators of this pathways is Ras, a GTPase that, upon activation, binds GTP to “turn on” the subsequent signaling cascade. Ras converting enzyme 1 (RCE1) post-translationally cleaves the 3 residues on the C-terminus of Ras, allowing Ras to properly localize to the plasma membrane. [35]

USP17 acts to deubiquitinate K63-ubiquitin domains on RCE1. [33] Such stabilization of RCE1 allows for proper localization of Ras, thus promoting proliferation upon activation of early receptors in the ERK Pathway. Ras hyperactivity can result in cell cycle dysregulation. [36] Thus, regulation of Ras through USP17 acts as another point in Ras regulation.

USP Regulation of G1-S Transition Edit

Cyclin-dependent kinases (CDKs) are a family of enzymes that phosphorylate serine and threonine residues to drive the cell through the cell cycle. Activation of CDK2 is critical for the G1-S transition. For CDK2 to be activated, cyclin A must bind to the cyclin-dependent kinase complex (CDKC). Cell division cycle 25A (CDC25A) is a phosphatase that removes an inhibitory phosphate group from CDK2. [37] While ubiquitination would mark CDC25A for degradation, thus blocking progression to S phase, USP17 deubiquitinates CDC25A. [33] An increase in CDC25A stability promotes CDKC activity, thus driving the cell through the G1-S transition.

USP17 also regulates cell cycle progression by acting on SETD8 to downregulate transcription of cyclin-dependent kinase inhibitor 1 (CDKN1A), also known as p21. [33] CDKN1A binds to and inhibits CDK2 using its N-terminal binding domain, thus blocking progression through the G1-S transition. SETD8, a methyltransferase, uses S-Adenosyl methionine to methylate the Lys20 residue of histone 4, resulting in the condensation of chromosomes. [38] This compaction of the DNA downregulates CDKN1A transcription. USP17 deubiquitinates SETD8, thus reducing its propensity for degradation and increasing its intracellular stability. [33] The resulting downregulation in CDKN1A transcription promotes CDK2 activity, allowing the cell to progress through the G1-S transition. See schematic of the role of DUBs in the cell cycle regulation. [33]

USP44 in Anaphase Initiation Edit

The spindle checkpoint (also referred to as the mitotic checkpoint) ensures proper separation of chromosomes. Broadly, the mitotic checkpoint promotes fidelity in chromosomal segregation, increasing the likelihood that each daughter cell receives only one duplicated chromosome. [39] Such a mechanism is crucial, as errors in chromosomal separation have been implicated in cancer, birth defects, and antibiotic resistance in pathogens. [40] One of the core regulator proteins is the anaphase-promoting complex (APC/C). APC/C ubiquitinates securin. [41] The resulting destruction of securing release separase, [39] which hydrolyzes cohesion – the protein that binds sister chromatids together.

New research from Stegmeier and colleagues [34] published in the journal Nature demonstrates a crucial role for USP44 in regulating the spindle checkpoint. Using an shRNA screen, USP44 was identified to stabilize the inhibition of APC/C [34] The binding of CDC20 to APC/C is required for the ubiquitination of securin. [42] A protein called hMAD2 can form an inactive trimer with APC and CDC20, forming the hMAD2-CDC-APC complex. [42] Upon the ubiquitination of CDC20 by UbcH10, hMAD2 dissociates, and APC/C becomes active. [43] It is important to note that ubiquitination of CDC20 does not serve to mark it for degradation, but rather promote dissociation of hMAD2 from the hMAD2-CDC-APC complex. USP44, a ubiquitin-specific-processing protease, can stabilize the inactive hMAD2-CDC-APC complex by counteracting UbcH10 ubiquitination. This blocks hMAD2 dissociation and allows for proper regulation of APC/C, keeping it inactive until proper attachment of the mitotic spindle. Upon proper attachment, switch-like behavior allows for the activation of APC/C. [34] This results in the cleavage of cohesion, allowing for the separation of sister chromatids.

DNA damage can prove catastrophic for an organism. Mechanisms for DNA mutation include oxidative stress, DNA replication errors, exogenous carcinogens, radiation, and spontaneous base mutation. Upon DNA damage, cell cycle progression is halted to prevent propagation of the mutation. The TP53 gene (also known as p53) is crucial in ensuring the conservation of the genome. [44] Deubiquitinating enzymes play an integral role in maintaining p53’s function.

In healthy cells, p53 activates the E3 ubiquitin ligase MDM2 which in turn ubiquitinates p53. This creates a negative feedback loop, whereby the degradation of p53 allows for cells to flow through the cell cycle. [45] Upon DNA damage, Ubiquitin-specific-processing protease 7 (USP7) stabilizes p53 by cleaving ubiquitin. [46] For USP7 to deubiquitinate p53, it must localize to the nucleus. However, no nuclear localization sequence (NLS) has been found. [47] Despite no known NLS, one study showed that, upon deletion of USP7’s N-terminus, no nuclear localization occurred. [47] It is possible that other proteins facilitate nuclear entry of USP7.

Once stabilized, p53 can exert its tumor suppression function. Downstream pathways of p53 act to either halt cell cycle progression in G1 or G2 phases of the cell cycle [48] or promote cell-death, depending on the severity of the DNA damage. [49] See schematic of the role of USP7 in the p53-dependent pathway. [48] or promote cell-death, depending on the severity of the DNA damage. [49] See schematic of the role of USP7 in the p53-dependent pathway. [49]


Stereochemical Assignment

The structure of cytostatin was disclosed without a definition of its relative or absolute stereochemistry, prompting us to secure a stereochemical assignment prior to initiating the synthetic work. In previous efforts, we defined the (5S,9S,11S)-stereochemistry for fostriecin, 10a and this assignment was extended to cytostatin based on its structural and functional similarity ( Figure 3a ). In these studies, we observed an intramolecular hydrogen bond between the C9-phosphate and C11-hydroxy group of fostriecin and demonstrated that the resulting cyclic structure exists in a rigid twist boat conformation ( Figure 3b ) that gives rise to distinct 1 H– 1 H coupling constants between H11 and H10a (syn) or H10b (anti) (J10a,11 = 3.7 Hz, J10b,11 = 9.6 Hz). Based on a well-founded assumption of a similar cyclic structure for cytostatin, its reported H10–H11 coupling constant (J10,11 = 9.4 Hz) 1a indicated a 10,11-anti configuration for the natural product. Alternatively, assumption of the adoption of one of two possible hydrogen bonded chair conformations for the C9� structure of cytostatin gives rise to a similar anti H10–H11 coupling constant and led Waldmann 14 to the same (10S)-stereochemical assignment ( Figure 3c ).

Stereochemical assignment for cytostatin.

To decipher the C4� relative stereochemistry, four diastereomeric lactones (C1� partial structures) were synthesized for spectral comparison ( Scheme 1 ). Construction of lactone 12 began with silylation (TBDPSCl, imidazole, DMF, 25 ଌ, 2 h, 99%) of methyl (S)-3-hydroxy-2-methylpropionate (7) followed by conversion of the ester to the corresponding aldehyde 9 (DIBAL-H, toluene, � ଌ, 1 h TPAP, NMO, CH2Cl2, 0 ଌ, 30 min, 82%, 2 steps). 21 Crotylation of 9 (cis-2-butenyldiisopinocampheylborane, THF, ether,� ଌ, 12 h NaOH, H2O2, 70 ଌ, 5 h, 63%, 8:1 dr) 22 gave alcohol 10 which was converted to the α,β-unsaturated lactone 12 via acylation (acryloyl chloride,i-Pr2NEt, CH2Cl2, 0 ଌ, 2 h, 91%) and subsequent ring closing metathesis (Grubbs' I catalyst, CH2Cl2, 40 ଌ, 12 h, 89%). 23 This sequence was used to access lactones 13, 14, and 15 in a stereodivergent manner by treating (R)- or (S)-9 with the appropriate crotylboration reagent followed by acylation and ring closing metathesis ( Scheme 1 ).

Synthesis of 1215.

Upon examination of the 1 H NMR spectra of 1215, it became apparent that the 4,5-syn and 4,5-anti configurations exhibit distinct coupling constants ( Figure 4 ). 24 Thus, the coupling constants observed for cytostatin were indicative of a 4,5-syn relative configuration. 25 Of the two 4,5-syn lactones, the 1 H NMR spectrum of 12 was closer to that of cytostatin as exemplified by an average absolute difference in chemical shift values (for H2–H6) of only 0.08 ppm, suggesting a 5,6-anti relative configuration for the natural product. Most notably, the chemical shifts (CD3OD) of 13 for H4, H5, 4-CH3 (δ 1.18) and especially the key 6-CH3 (δ 0.88) were considerably distinct from those of 1 (4-CH3 at δ 1.00, 6-CH3 at δ 0.98) and 12 (4-CH3 at δ 1.08, 6-CH3 at δ 0.99). Based on this analysis, the (4S,5S,6S,9S,10S,11S) configuration was assigned to cytostatin. Independent of our efforts, Waldmann 14 conducted and has disclosed an analogous set of studies using a different lactone series and a similar retrospective interpretation of the cytostatin spectroscopic properties to arrive at the same stereochemical assignments for which our relative and absolute stereochemical assignments of the fostriecin distal C4 and C9/C11 centers served as the foundation. 10

1 H NMR comparison for lactones 1215.

In digesting the conformational features embedded in this segment of cytostatin, we came to recognize that the distinguishing H5–H6 coupling constant was also diagnostic of the side chain adopting a single, preferred conformation ( Figure 4b ). This single side chain orientation is adopted to avoid the syn pentane interactions of the two alternatives (ΔE ca. 3.7 kcal/mol). In addition to accounting for the large H5–H6 coupling constant (J = 9.5�.2 Hz) characteristic of their anti relationship in this conformation, this also suggests that the role of the cytostatin C4 and C6 methyl groups is to confine the side chain to a single orientation facilitating binding to PP2A, a potential role that is reinforced in our retrospective modeling of the cytostatin binding to a PP2A homology model.

Synthetic Plan

We designed a synthetic route to cytostatin that would confirm its stereochemical assignment and provide rapid access to the C10� stereoisomers of 1 that could be used to define the role of the C11-hydroxy and C10-methyl group in PP2A binding. The convergent approach relied on installation of the triene at a late stage and in a single-step by addition of 18 to a C11 aldehyde, enlisting substrate control to set the C11 stereochemistry. Assembly of the C7� bond by coupling a cuprate derived from iodide 16 with epoxide 17 isolates the two stereotriads of cytostatin, allowing for independent adjustment of their stereochemistry. In addition, this bond construction provided the opportunity to utilize lactones 1215 as intermediates in the total synthesis of the natural product and its C4� diastereomers. A Sharpless epoxidation served to set the C9 and C10 stereochemistry, and the triene was synthesized in a short, stereospecific approach relying on an electrocyclic ring opening to set the geometry of the three olefins ( Figure 5 ).

Synthetic plan for cytostatin.

Synthesis of C1�

Lactone 12, developed during our work to secure a stereochemical assignment for cytostatin, served as the starting point for synthesis of the C1� fragment of cytostatin ( Scheme 2 ). The electrophilic α,β-unsaturated lactone was masked by carbonyl reduction (DIBAL-H, CH2Cl2, � ଌ, 30 min) and conversion of the intermediate lactol to the methyl acetal 19 (PPTS, MeOH, 25 ଌ, 10 min, 82%, 2 steps). Acetal 19 formed as a single isomer 26 under these conditions, but equilibrated to a 2:1 mixture of anomers when subjected to MeOH/PPTS treatment for longer reaction times. Although the C1 stereocenter would eventually be removed, selective formation of (1R)-19 allowed us to carry forward a single diastereomer, simplifying the isolation and characterization of intermediates in the synthesis. Interestingly, the alternative and more direct generation of 19 employing the corresponding methoxy acetal precursor in the Grubbs' metathesis cyclization proved less satisfactory with the substrate being more challenging to effectively prepare from 10 (30�%) and the product being a mixture of C1 diastereomers. After desilylation of 19 (Bu4NF, THF, 25 ଌ, 12 h) to give 20, efforts to directly convert the alcohol to the corresponding halide proved problematic as 20 either remained unchanged (PPh3, NIS PPh3, CBr4) or decomposed (PPh3, I2) under standard conditions. However, 20 could be converted to iodide 16 through a 2-step process of tosylation (p-TsCl, NaH, benzene, 25 ଌ, 4 h, 81%, 2 steps) followed by iodide displacement (NaI, acetone, 56 ଌ, 12 h, 90%). The C1� precursor 16 contains the C4� stereotriad of cytostatin, installed in a stereochemically-tunable sequence through Brown crotylboration of chiral building block 9.

Synthesis of C8�

Assembly of the C8� epoxide unit ( Scheme 3 ) began with 22, which was prepared on a multigram scale by a known procedure 27 that relies on a Sharpless asymmetric epoxidation reaction to set the chirality of the epoxy alcohol. Epoxide opening using a procedure developed by Tius (MeMgI, CuI, Et2O, THF, � ଌ, 3 h, 69%) 28 provided a 3:1 mixture of 23 and its regioisomer (1,2-diol), which was removed by periodate treatment (NaIO4, H2O, Et2O, 25 ଌ, 1.5 h, 91%) followed by chromatography. After debenzylation (H2, Pd/C, MeOH, 25 ଌ, 2h, 99%), triol 24 29 was transformed to epoxide 17 in a sequence that mirrors a reported conversion of epi-(3R)-24 into its corresponding epoxide. 30 Thus, 1,2-diol protection (3,3-dimethoxypentane, p-TsOH, DMF, 25 ଌ, 12 h, 68%) followed by PMB ether formation (NaH, PMBCl, Bu4NI, THF, 0 to 20 ଌ , 20 h, 80%) gave 26. After acetal removal (p-TsOH, MeOH, 25 ଌ, 7 h, 89%), diol 27 was transformed to epoxide 17 through tosylation of the primary alcohol followed by its intramolecular displacement (NaH, pTs-imidazole, THF, � to 25 ଌ, 4 h, 82%). The C8� unit bears the C9 and C10 stereocenters of cytostatin that contain the subtituents proposed to affect PP2A active site binding. Alternative C9� stereochemical configurations of cytostatin are accessible by transforming the stereoisomers of triol 24 to their corresponding epoxides through this simple and reliable synthesis.

Synthesis of C12�.

Crucial to the implementation of the convergent synthetic plan set forth herein was the ability to supply and successfully manipulate the potentially labile bromotriene 18. In a modification of Taylor's synthesis of Z,E-dienals, 31 addition of MeLi (THF, � ଌ, 2 h) to pyrylium tetrafluoroborate followed by electrocyclic ring opening provided 28 in 66% as a single isomer ( Scheme 4 ). Transformation of 28 to dibromoolefin 29 (CBr4, PPh3, Et3N, CH2Cl2, 0 ଌ, 10 min, 97%) 32 and selective (E)-bromide reduction (Bu3SnH, Pd(PPh3)4, ether, 0 ଌ, 15 min, 95% conversion) 33 gave 18 stereoselectively. Isolation of bromide 18 was complicated by its volatility and tendency to polymerize during distillation or concentration. However, chromatographic purification (silica gel, pentane) followed by careful concentration in the presence of Et3N to prevent trace acid-induced polymerization allowed isolation of 18 in superb yields (73%) considering its lability. Storage of bromide 18 proved more problematic, but it was readily generated immediately prior to use from the dibromide 29 which was stable for months when stored as a refrigerated benzene solution. The C12� precursor 18 contains the (Z,Z,E)-conjugated triene, ready for incorporation into cytostatin as a single unit.

Synthesis of Cytostatin.

Assembly of 1 ( Scheme 5 ) was initiated by converting iodide 16 into the corresponding cuprate (t-BuLi, ether, � ଌ, 5 min then (2-Th)CuCNLi, THF, � to 0 ଌ, 5 min) 34 followed by addition of 17 at 0 ଌ to give 30 in 84%. The coupling of 16 and 17 was slow when alternative metalated forms of 16 were used (i.e. R2CuLi, RMgI/cat. CuI, R2CuCNLi2, RLi/BF3), and the success of the transformation was dependent on the use of the higher order cuprate at 0 ଌ and at concentrations of 50 mM or greater. Acetal formation (ethyl vinyl ether, PPTS, CH2Cl2, 25 ଌ, 30 min, 90%) gave 31 as a 1.2:1 mixture of diastereomers which, for ease of isolation and characterization, were separated after PMB removal (DDQ, CH2Cl2, H2O, 25 ଌ, 30 min, 85%) and carried through the subsequent 4 steps of the synthesis separately. Oxidation of 32 (DMP, CH2Cl2, 25 ଌ, 15 min, 91%) 35 provided aldehyde 33 setting the stage for installation of the triene. Bromine/lithium exchange on 18 followed by addition of 33 gave 34 as a 2:1 mixture of diastereomers favoring the undesired 10,11-syn isomer (Felkin product). Following Still's precedent, 36 the diastereoselectivity of the addition could be modulated by using a copper-based nucleophile, which promotes addition to a chelated β-alkoxy aldehyde. Thus, conversion of 18 to its Bu3P-stabilized cuprate (t-BuLi, ether, � ଌ, 1.5 h CuI–PBu3, 10 min) prior to addition of 33 (� ଌ, 30 min, 76%) produced 34 as a pair of chromatographically-separable diastereomers in a ratio of 7:1 favoring the desired 10,11-anti product. As anticipated, the diastereoselectivity of the triene addition was also highly dependent on the C9 protecting group ( Figure 6 ). When the non-chelating TES group was used instead of EE, the Felkin product (syn) predominated when either the organolithium or organocuprate nucleophile was added to the aldehyde. This observation is consistent with the Felkin versus chelation model for the diastereoselective addition and proved useful for the synthesis of the cytostatin stereoisomers. Assignment of the C10� relative stereochemistry of anti-34 was initially based on its H10–H11 coupling constant (J = 9.0 Hz), which was indicative of a 10,11-anti configuration and consistent with a C9-alkoxy/C11-hydroxy hydrogen bonded cyclic structure ( Figure 3 ), and this stereochemical assignment was later confirmed through analysis of the 9,11-acetonide derived from a more advanced synthetic intermediate. The minor isomer (syn-34) from the β-chelation controlled triene addition exhibited an H10–H11 coupling (J = 3.6 Hz) expected to arise from a 10,11-syn configuration.

Diastereoselectivity of the triene addition.

Following silylation (TBSCl, imidazole, DMF, 25 ଌ, 2 h, 95%) of alcohol 34, treatment of 35 with dilute HCl for short reaction times (0.02 N HCl, acetone, water, 25 ଌ, 10 min, 85%) induced simultaneous C1 and C9 acetal hydrolysis without affecting the silyl ether, whereas more conventional acidic deprotection methods (0.2 N HCl�tone 80% aq. HOAc 90% TFA–H2O) led to competitive global deprotection even when conducted at low temperatures. In this context, the use of the C9 EE protecting group proved valuable as it served not only to direct the formation of the C11 stereocenter (through chelation), but could be effectively removed in the presence of the labile silyl ether and conjugated triene. Selective oxidation (Ag2CO3�lite, benzene, 80 ଌ, 2 h, 80%) of lactol 36 produced 37, which was phosphorylated (i-Pr2NP(OFm)2, tetrazole, CH3CN, CH2Cl2, 25 ଌ, 15 min H2O2, 10 min, 82%) using the protocol introduced by Waldmann 14c to give 38. Desilylation (HF–pyr, THF, pyridine, 25 ଌ, 4 h, 85%) followed by fluorenylmethyl removal (Et3N, CH3CN, 25 ଌ, 17 h, 99%) provided 1, identical to a sample of natural cytostatin ( 1 H NMR, TLC, HPLC, HRMS).

At this stage, the C9� relative stereochemistry of synthetic cytostatin was confirmed by examining the spectral properties of acetonide 41, derived from intermediate 37. Cleavage of the silyl ether of37 (HF–pyr, THF, pyridine, 25 ଌ, 2 h, 99%) provided dephosphocytostatin (40), which was subjected to acetal formation (2,2-dimethoxypropane, p-TsOH, THF, 25 ଌ, 1 h, 99%) to deliver 41 in excellent yield. The acetonide methyl groups (δ 24.6, 25.1) and ketal carbon (δ 101.2) of 41 exhibited 13C NMR chemical shifts (CD3CN, 150 MHz) characteristic of anti-1,3-diol acetonides (δ 23.6�.6 and 100.2�.0) and distinct from those observed for syn-1,3-diol acetonides (δ 18.6�.9/29.8�.2, and 98.0�.3). 37 Furthermore, the H10–H11 coupling constant (J = 8.4 Hz) confirmed the expected 10,11-anti stereochemistry. Dephosphocytostatin (40) displayed spectral properties similar to 41, and particularly striking is its H10–H11 coupling constant (J = 8.4 Hz) and C10-methyl group 13 C NMR chemical shift (δ 10.8, CD3CN, 150 MHz), both of which are indistinguishable from those of 41. This comparison suggests that the C9� hydrogen-bonded cyclic structure of 40 (and 1) exists in the twist boat conformation observed for 41 and fostriecin ( Figure 3b ) rather than the alternative chair conformation ( Figure 3c ) suggested by Waldmann. The latter places the C10-methyl group in an equatorial position that would result in a downfield shifted 13 C NMR resonance for the C10-Me instead of the identical signal that is observed.

Synthesis of Cytostatin Diastereomers

The completion of the total synthesis of cytostatin set the stage for preparation of the C10� diastereomers (4244, Figure 8 ) that were used to confirm the C9� stereochemical assignments and to further delineate the cytostatin–PP2A interaction. epi-(11R)-Cytostatin (42) was synthesized by advancing the 10,11-syn isomer of 34 through the final 6 steps of the cytostatin synthesis, which occurred without incident and was conducted without significant optimization of the individual steps ( Scheme 6 ). As noted above, syn-34 was the major product of the addition of triene 18 to aldehyde 33 when the lithium deriviative of 18 was used as the nucleophile ( Figure 6 ), and this procedural adjustment allowed accumulation of sufficient material to produce 42.

Structures of cytostatin C10� diastereomers.

Synthesis of 42.

The synthesis of cytostatin isomers 43 and 44 bearing the (10R) configuration was achieved by utilizing the known epoxide 50 30 in place of 17 in the synthesis ( Scheme 7 ). A key element of the derivation of the cytostatin synthesis in which the two stereotriads are independently created and joined in a convergent epoxide opening followed by a single-step, late stage addition of the intact triene with diastereocontrolled introduction of the C11 center is that the approach provides rapid, divergent access (11 steps from convergence point) to each C9� diastereomer. Thus, coupling of 16 and 50 proceeded as expected under the previously optimized conditions (t-BuLi, ether, � ଌ, 5 min then (2-Th)CuCNLi, THF, � to 0 ଌ, 5 min 50, ether, 0 ଌ, 1 h, 72%) to provide 51 in superb yield. A C9 TES group was selected in preference to EE for synthesis of the (10R) series since β-chelation was not required for installation of the natural (11S)-configuration and the silyl ether formation does not produce additional diastereomers. Thus, through a sequence of silylation (TESCl, imidazole, DMF, 25 ଌ, 1 h, 87%), PMB removal (DDQ, CH2Cl2, H2O, 25 ଌ, 15 min, 72%), and oxidation (DMP, CH2Cl2, 25 ଌ, 10 min, 90%), 35 51 was converted to aldehyde 54. Addition of 54 to the lithium derivative of 18 (t-BuLi, ether, � ଌ, 1.5 h 54, � ଌ, 1 h, 70%) produced 55 as a 2:1 mixture of diastereomers favoring the Felkin 10,11-syn product ( Figure 6 ). The expected stereochemical assignments were confirmed based on the H10–H11 coupling for 55 (J10,11 = 9.0 for anti, J10,11 = 2.4 for syn). After chromatographic separation, syn-55 and anti-55 were converted to 43 and 44 by way of the process developed for the synthesis of 1 ( Scheme 7 ). Of note, the TES group was also cleaved at a useful rate under the conditions (0.02 N HCl, acetone, water, 25 ଌ, 10 min, 61%) developed for removal of the EE group in the presence of the triene and labile allylic TBS ether. Thus, the utility of our convergent total synthesis of cytostatin is highlighted by the fact that only 23 additional steps were required to prepare its entire complement of C10� diastereomers.

Synthesis of 43 and 44.

Confirmation of Stereochemical Assignment

The stereochemistry of the C9� portion of cytostatin has thus far been assigned based on a key coupling constant (J10,11 = 9.4 Hz) that suggests a 10,11-anti relative stereochemistry and the assumption that cytostatin and fostriecin have identical absolute configurations at shared chiral centers (C9 and C11). While this is sound reasoning and synthetic 1 proved spectroscopically indistinguishable from natural cytostatin, 14 , 20 the magnitude of the optical rotation of synthetic 1 reported both by Waldmann 14c ([α] 20 D +46 (c 0.27, MeOH)) and found herein ([α] 25 D +45 (c 0.07, MeOH)) did not match that measured for natural cytostatin ([α] 25 D +20 (c 0.27, MeOH)) by Waldmann on an aged authentic sample, 14c leaving room for doubt about the identity of natural cytostatin. Visual comparison of the 1 H NMR spectra of the four C10� diastereomers (1, 42, 43, 44, Figure 9 ) with that of natural cytostatin demonstrates that synthetic 1 matches natural cytostatin identically, while 4244 display substantially different resonances arising from C11-H, C9-H, and C10-Me. In general, the C9 signals for the 10,11-syn isomers (42, 43) were further upfield than for the 10,11-anti isomers (1, 44), while the C11 signals in the syn compounds were further downfield than those of the anti isomers. The 10,11-anti isomer 44 most closely resembles 1 spectrally, but the large difference in the shift of their H11 protons (Δδ = 0.22 ppm) clearly distinguishes the two. The sensitivity of the 1 H NMR signals to stereochemical changes in the C9� region and the magnitude of the H10–H11 coupling for each diastereomer can be best interpreted by their adoption of a C9-phosphate/C11-hydroxy hydrogen bonded twist boat conformation. These results also remove any doubt about the 9,10-relative configuration since similarly large spectral changes would be observed if the C9� cyclic structure were altered through that locus. In total, the dissonance of the alternative C10� diastereomers with cytostatin, the comparison of the C1� lactone partial structures with cytostatin, the spectral identity of synthetic 1 with natural cytostatin, the biological profile of synthetic 1, and the fact that synthetic 1 shares the same sign of optical rotation with natural cytostatin authenticate the (4S, 5S, 6S, 9S, 10S, 11S) assignment. The discrepancy in the magnitude of the optical rotation of synthetic 1 and natural cytostatin has been attributed by Waldmann to impurities in his natural sample that resulted from long-term storage. 14c

Partial 1 H NMR comparison for natural cytostatin, 1, and 4244.

Synthesis of Additional Key Analogues

In addition to the C10� diastereomers, our synthetic route provided access to several other analogues useful for defining the impact of the C9� substituents and the triene unit on PP2A inhibition. Intermediate 37, ideally suited for the preparation of phosphate replacements, provided not only dephosphocytostatin (40) but by sulfation (SO3–pyridine, THF, 25 ଌ, 30 min, 71%) followed by desilylation (HF–pyr, THF, pyridine, 25 ଌ, 1.5 h, 83%) also provided 67, the sulfated version of cytostatin ( Scheme 8 ).

Synthesis of cytostatin analogues.

Conversion of acetal 30 to its corresponding lactone through hydrolysis (0.02 N HCl, acetone, water, 25 ଌ, 10 min, 99%) and oxidation (Ag2CO3�lite, benzene, 80 ଌ, 2 h, 96%) gave 68. Phosphorylation (i-Pr2NP(OFm)2, tetrazole, CH3CN, CH2Cl2, 25 ଌ, 15 min H2O2, 10 min, 88%) of 68 was followed by Et3N treatment (Et3N, CH3CN, 25 ଌ, 20 h, 88%) to provide the phosphate 70 ( Scheme 8 ). Alternatively, cleavage of the PMB group of 69 (DDQ, CH2Cl2, H2O, 25 ଌ, 30 min, 70%) followed by Et3N treatment (Et3N, CH3CN, 25 ଌ, 17 h, 99%) produced 72, a cytostatin C1� partial structure that lacks the C12� triene unit.

To provide a means to quantify the impact of the C11-hydroxy on PP2A binding, 11-deshydroxy cytostatin (78) was constructed through a late-stage modification of the cytostatin total synthesis ( Scheme 9 ). Alcohol 32 was transformed to iodide 73 (NIS, PPh3, imidazole, THF, 25 ଌ, 5 min, 74%), a suitable coupling partner for bromotriene 18 at the correct oxidation state to deliver 78. The union of 18 and 73 was achieved through alkylation of the cuprate derived from 18 (t-BuLi, ether, � ଌ, 1 h CuI–PBu3, 10 min 73, 0 ଌ, 1 h, 63%) to provide 74 in good yield with only minimal efforts at optimization. Notably, this cuprate coupling represents a nice example of a sp 2 -sp 3 organometallic coupling reaction enlisting an unactivated sp 3 halide complementary to the more contemporary Stille, Suzuki, and Negishi reactions of vinyl stannanes, boronic acids, or organic zinc reagents currently being explored with sp 3 halides. Acetal hydrolysis (0.02 N HCl, acetone, water, 25 ଌ, 10 min, 77%) followed by lactol oxidation (Ag2CO3�lite, benzene, 80 ଌ, 30 min, 76%) gave lactone 76, which after phosphorylation (i-Pr2NP(OFm)2, tetrazole, CH2Cl2, CH3CN, 25 ଌ, 30 min H2O2, 5 min, 65%) and Et3N treatment (Et3N, CH3CN, 25 ଌ, 17 h, 79%) provided 78.

Synthesis of 78.

Total synthesis and evaluation of cytostatin, its C10-C11 diastereomers, and additional key analogues: impact on PP2A inhibition

The total synthesis of cytostatin, an antitumor agent belonging to the fostriecin family of natural products, is described in full detail. The convergent approach relied on a key epoxide-opening reaction to join the two stereotriad units and a single-step late-stage stereoselective installation of the sensitive (Z,Z,E)-triene through a beta-chelation-controlled nucleophilic addition. The synthetic route provided rapid access to the C4-C6 stereoisomers of the cytostatin lactone, which were prepared and used to define the C4-C6 relative stereochemistry of the natural product. In addition to the natural product, each of the C10-C11 diastereomers of cytostatin was divergently prepared (11 steps from key convergence step) by this route and used to unequivocally confirm the relative and absolute stereochemistry of cytostatin. Each of the cytostatin diastereomers exhibited a reduced activity toward inhibition of PP2A (>100-fold), demonstrating the importance of the presence and stereochemistry of the C10-methyl and C11-hydroxy groups for potent PP2A inhibition. Extensions of the studies provided dephosphocytostatin, sulfocytostatin (a key analogue related to the natural product sultriecin), 11-deshydroxycytostatin, and an analogue lacking the entire C12-C18 (Z,Z,E)-triene segment, which were used to define the magnitude of the C9-phosphate (>4000-fold), C11-alcohol (250-fold), and triene (220-fold) contribution to PP2A inhibition. A model of cytostatin bound to the active site of PP2A is presented, compared to that of fostriecin, which is also presented in detail for the first time, and used to provide insights into the role of the key substituents. Notably, the alpha,beta unsaturated lactone of cytostatin, like that of fostriecin, is projected to serve as a key electrophile, providing a covalent adduct with Cys269 unique to PP2A, contributing to its potency (> or =200-fold for fostriecin) and accounting for its selectivity.

Synthetic Biology and Metabolic Engineering in Plants and Microbes Part B: Metabolism in Plants

E.C. O’Neill , . R.A. Field , in Methods in Enzymology , 2016

2.2 Nonribosomal Peptide Synthetases

NRPs are a range of small molecules with a diversity of functions that include siderophores such as enterobactin ( Walsh, Liu, Rusnak, & Sakaitani, 1990 ), antibiotics such as actinomycin ( Schauwecker, Pfennig, Schröder, & Keller, 1998 ), and toxins such as microcystin ( Noguchi et al., 2009 ). NRPs are formed by the sequential condensation of amino acids, including both proteinogenic and nonproteinogenic examples. They are synthesized by nonribosomal peptide synthetases (NRPSs), multimodular megasynthases: each module comprises an adenylation domain (A), which selects the incoming amino acid, activates it by adenylation, and attaches it to a peptidyl carrier protein (PCP) via a thiol-containing phosphopantetheine arm, and a condensation domain (C) which catalyzes the formation of an amide linkage in the growing peptide (see Fig. 2 ) ( Walsh, 2016 ). As for PKSs, there is a TE domain which catalyzes the off-loading of the completed peptide and there can be accessory domains within the megasynthases, encodings N-methyltransferases or amino acid epimerases, for instance. The amino acid specificity of the A domain can be predicted with reasonable certainty based on the amino acid residues that line the binding pocket of this domain ( Stachelhaus, Mootz, & Marahiel, 1999 ). NRPs can be modified by, for example, initiation with a lipid chain or formation of an ester between side chains and the C-terminus to cyclize the product, as is seen in arthrofactin, a member of the large class of cyclic lipopetide antibiotics ( Roongsawang et al., 2003 ). There are very few examples of bioactive peptides that have been purified from eukaryotic algae: those that have been, such as the galaxamide from the red algal Galaxaura ( Xu et al., 2008 ) and kahalalide F from the green alga Bryopsis ( Hamann & Scheuer, 1993 ), were purified from macroalgae, collected from the wild, and so a bacterial origin for these compounds cannot be ruled out. There is no data on the biosynthesis of these compounds, thought their structure, including, N-methylation in the former and d -amino acids in the latter, and cyclization suggests an NRPS origin for these compounds.

As both PKSs and NRPSs have modular and linear biosynthetic logic, hybrid arrangements can occur in which an amino acid can be extended with a ketide or an amino acid incorporated into a PK. For example, in the former case the proteasome inhibitor epoxomicin has a peptide backbone extended by a ketide to form the epoxide warhead, which forms a covalent adduct with its’ target enzyme ( Schorn et al., 2013 ). Curacin is an example of the latter, in which a cysteine residue is added into a growing PK and then decarboxylated to continue the chain extension ( Chang et al., 2004 ).

Title: PNT1 is a C11 cysteine peptidase essential for replication of the Trypanosome Kinetoplast

The structure of a C11 peptidase PmC11 from the gut bacterium, Parabacteroides merdae, has recently been determined, enabling the identification and characterization of a C11 orthologue, PNT1, in the parasitic protozoon Trypanosoma brucei. A phylogenetic analysis identified PmC11 orthologues in bacteria, archaea, Chromerids, Coccidia, and Kinetoplastida, the latter being the most divergent. A primary sequence alignment of PNT1 with clostripain and PmC11 revealed the position of the characteristic His-Cys catalytic dyad (His 99 and Cys 136 ), and an Asp (Asp 134 ) in the potential S1 binding site. Immunofluorescence and cryoelectron microscopy revealed that PNT1 localizes to the kinetoplast, an organelle containing the mitochondrial genome of the parasite (kDNA), with an accumulation of the protein at or near the antipodal sites. Depletion of PNT1 by RNAi in the T. brucei bloodstream form was lethal both in in vitro culture and in vivo in mice and the induced population accumulated cells lacking a kinetoplast. In contrast, overexpression of PNT1 led to cells having mislocated kinetoplasts. RNAi depletion of PNT1 in a kDNA independent cell line resulted in kinetoplast loss but was viable, indicating that PNT1 is required exclusively for kinetoplast maintenance. Expression of a recoded wild-type PNT1 allele, but not of an active site mutantmore » restored parasite viability after induction in vitro and in vivo confirming that the peptidase activity of PNT1 is essential for parasite survival. Furthermore, these data provide evidence that PNT1 is a cysteine peptidase that is required exclusively for maintenance of the trypanosome kinetoplast. « less

Protein kinases and phosphatases involved in the acclimation of the photosynthetic apparatus to a changing light environment

Photosynthetic organisms are subjected to frequent changes in light quality and quantity and need to respond accordingly. These acclimatory processes are mediated to a large extent through thylakoid protein phosphorylation. Recently, two major thylakoid protein kinases have been identified and characterized. The Stt7/STN7 kinase is mainly involved in the phosphorylation of the LHCII antenna proteins and is required for state transitions. It is firmly associated with the cytochrome b6f complex, and its activity is regulated by the redox state of the plastoquinone pool. The other kinase, Stl1/STN8, is responsible for the phosphorylation of the PSII core proteins. Using a reverse genetics approach, we have recently identified the chloroplast PPH1/TAP38 and PBPC protein phosphatases, which counteract the activity of STN7 and STN8 kinases, respectively. They belong to the PP2C-type phosphatase family and are conserved in land plants and algae. The picture that emerges from these studies is that of a complex regulatory network of chloroplast protein kinases and phosphatases that is involved in light acclimation, in maintenance of the plastoquinone redox poise under fluctuating light and in the adjustment to metabolic needs.

1. Introduction

Photosynthetic organisms have the remarkable ability to adapt rapidly to changes in light conditions. This is particularly true for plants and algae, which have developed specific mechanisms that allow them to act either as energy dissipators (when the light energy absorbed by the light-harvesting system exceeds the capacity of the photosynthetic system) or as energy collectors (when the absorbed light is limiting). In these organisms, the primary reactions of photosynthesis occur in the thylakoid membranes and are catalyzed by the photosynthetic complexes photosystem II (PSII), the cytochrome b6f complex (cyt b6f) and photosystem I (PSI). These three complexes form the photosynthetic electron transport chain in which two serially connected photochemical reactions in PSII and PSI lead to the oxidation of water, followed by electron flow through the plastoquinone pool, cyt b6f and plastocyanin, and ultimately to the reduction of ferredoxin. These processes are responsible for the formation of both reducing power and a proton gradient across the thylakoid membrane that is used to produce ATP through the ATP synthase complex. Thylakoid membranes are compartmentalized in grana regions consisting of appressed membranes and stromal lamellae that often connect grana stacks to one another. It has been known since the pioneering studies of Andersson & Andersson [1] that while PSII and LHCII (light-harvesting complex II) are mostly confined to the grana regions, PSI and ATP synthase are localized exclusively in the stroma lamellae because of their stromal domains, which prevent these complexes from entering the grana [2]. Cyt b6f is distributed equally between these two thylakoid domains.

When plants and algae are subjected to an irradiance that is in excess of that which can be used by photosynthesis, a large proton gradient is generated across the thylakoid membrane, which leads to the dissipation of the excess absorbed light energy into heat through non-photochemical quenching (for review see [3]). As an unavoidable consequence of its photochemical activity that generates one of the most oxidizing reactions observed in biological systems, PSII is damaged mainly at the level of the D1 reaction centre protein and needs to be repaired. Current evidence favours a model in which the damaged PSII moves out of the grana to the stromal lamellae where the damaged D1 protein is degraded through the concerted action of the FtsH and Deg proteases [4–6]. A newly synthesized D1 protein is then inserted into the PSII complex, which moves back to the grana region. This repair cycle requires a series of phosphorylation and dephosphorylation events at the level of the PSII core proteins D1, D2, CP43 and PsbH [7,8].

By contrast, when the light is limiting for growth, the photosynthetic machinery optimizes light capture and photosynthetic yield. Because the light-harvesting systems of PSII and PSI have distinct sizes and light absorption properties, their relative performance in light capture depends on both light quality and quantity. In particular, red and far-red light are preferentially absorbed by PSII and PSI, respectively. Thus, changes in the red/far-red ratio that occur under a canopy or through shading affect the relative yields of PSII and PSI. Under conditions that promote a preferential excitation of PSII relative to PSI, the redox state of the plastoquinone pool is more reduced. Binding of plastoquinol to the Qo site of the cyt b6f complex leads to the activation of a protein kinase that phosphorylates LHCII, the light-harvesting system of PSII. Phosphorylated LHCII (P-LHCII) dissociates from PSII and is displaced to PSI, where it increases the cross-section of the antenna system of PSI and thereby re-equilibrates the absorbed light between PSII and PSI. This process (called state transition) is reversible, as preferential excitation of PSI leads to the oxidation of the plastoquinone pool, inactivation of the kinase followed by dephosphorylation of LHCII, which moves back to PSII in the grana region (figure 1 for review, see [9–11]). This state corresponds to state 1, whereas the state in which P-LHCII is associated with PSI corresponds to state 2. State transitions were discovered independently over 40 years ago by Bonaventura & Myers [12] and Murata [13].

Figure 1. Scheme of state transitions. State transitions involve a redistribution of the mobile light-harvesting antenna of PSII (LHCII) between PSII and PSI. Upon preferential excitation of PSII, the plastoquinone pool (PQ) is reduced and plastoquinol docks to the Qo site of cyt b6f. This leads to the activation of a protein kinase that phosphorylates LHCII. The latter dissociates from PSII and migrates to PSI. If PSI is preferentially excited, the plastoquionone pool is oxidized, the kinase is inactivated and a phosphatase dephosphorylates the mobile LHCII, which moves back to PSII. (a) State 1 and (b) state 2 refer to the states in which the mobile LHCII is associated with PSII and PSI, respectively. Fd, ferredoxin PC, plastocyanin. Reproduced with permission from Rochaix [9].

It is interesting to note that while the mobile LHCII antenna in land plants constitutes only 15–20% of LHCII, in the green alga Chlamydomonas reinhardtii, it is considerably larger and represents 80 per cent of LHCII [14]. In this alga, transition to state 2 is associated with a switch from linear to cyclic electron flow and it occurs readily under conditions leading to a decrease in the cellular ATP level, such as anaerobic growth conditions or treatment of the cells with an uncoupler [15–17]. However, the difference between algae and plants may only be quantitative, as in all cases, LHCII phosphorylation responds to the redox state of the plastoquinone pool, which in turn is influenced by the metabolic state and the environmental conditions (see figure 4).Thus, it appears that an important role of state transitions in algae and plants is to maintain the redox poise of the plastoquinone pool and to respond to metabolic needs [11,18,19].

2. Genetic dissection of state transitions

Although phosphorylation of LHCII had already been demonstrated by Bennett [20] in the seventies, the biochemical hunt for the protein kinase involved in this phosphorylation was not successful in spite of numerous attempts [21–24]. We and others therefore decided to use a genetic approach in Chlamydomonas, on the basis of the observation that a large fraction of the LHCII antenna is mobile during state transitions, resulting in a large decrease in PSII fluorescence after a transition from state 1 to state 2 [25,26]. This property was used in a screen for mutants deficient in state transitions with a fluorescence video imaging system. Several mutants were isolated in this screen and are listed in table 1. One mutant of particular interest, stt7-7, was deficient in a protein kinase called Stt7 [27]. Besides being unable to perform state transitions, this mutant was deficient in LHCII phosphorylation under conditions favouring state 2 and blocked in state 1. Subsequently, two additional allelic mutants (called stt7-6 and stt7-9) were isolated. Among the mutants identified, some affect state transitions indirectly. This is the case for stt10 and stt2, which are deficient in the expression of MenC and MenD, two genes involved in the phylloquinone biosynthetic pathway [28]. In at least one of these mutants, phylloquinone, which normally acts as an electron carrier within PSI, is replaced by plastoquinone. This replacement leads to decreased performance of PSI and is the cause for the state transition phenotype. Two other mutants that have been partially characterized are C11 and HCM, which are locked in state 1 (S. Miras, S. Lobréaux and J.-D. Rochaix 2008, unpublished results). Because these mutants are leaky, it has not yet been possible to identify the genes responsible for the mutant phenotype.

The emergence of Mycobacterium tuberculosis (Mtb) resistance is a serious threat to public health. However, the quest for more efficient drugs against Mtb is hampered by the lack of a detailed understanding of Mtb virulence protein effectors. Here, we describe the swift modification of select Cys residues in multi-Cys proteins directly through chemistry. New insights into the biochemistry of emerging bacterial drug targets were obtained. We reveal a water Cys-Cys bridging mechanism that offers an explanation for the known resistance of Mtb protein tyrosine phosphatase A (PtpA) to the oxidative conditions that prevail within an infected host macrophage. This water Cys-Cys bridge motif is also found in the phosphatase SptpA from Staphylococcus aureus, suggesting its potential conserved structural role. The rationalization of the unique features of PtpA, an important target for Mtb drug discovery, could now be used in the design of novel small-molecule modulators.

The emergence of multidrug-resistant Mycobacterium tuberculosis (Mtb) strains highlights the need to develop more efficacious and potent drugs. However, this goal is dependent on a comprehensive understanding of Mtb virulence protein effectors at the molecular level. Here, we used a post-expression cysteine (Cys)-to-dehydrolanine (Dha) chemical editing strategy to identify a water-mediated motif that modulates accessibility of the protein tyrosine phosphatase A (PtpA) catalytic pocket. Importantly, this water-mediated Cys-Cys non-covalent motif is also present in the phosphatase SptpA from Staphylococcus aureus, which suggests a potentially preserved structural feature among bacterial tyrosine phosphatases. The identification of this structural water provides insight into the known resistance of Mtb PtpA to the oxidative conditions that prevail within an infected host macrophage. This strategy could be applied to extend the understanding of the dynamics and function(s) of proteins in their native state and ultimately aid in the design of small-molecule modulators.


The fold of fructose-1,6-bisphosphatase from pigs was noted to be identical to that of inositol-1-phosphatase (IMPase). [3] Inositol polyphosphate 1-phosphatase (IPPase), IMPase and FBPase share a sequence motif (Asp-Pro-Ile/Leu-Asp-Gly/Ser-Thr/Ser) which has been shown to bind metal ions and participate in catalysis. This motif is also found in the distantly-related fungal, bacterial and yeast IMPase homologues. It has been suggested that these proteins define an ancient structurally conserved family involved in diverse metabolic pathways, including inositol signalling, gluconeogenesis, sulphate assimilation and possibly quinone metabolism. [4]

Three different groups of FBPases have been identified in eukaryotes and bacteria (FBPase I-III). [5] None of these groups have been found in archaea so far, though a new group of FBPases (FBPase IV) which also show inositol monophosphatase activity has recently been identified in archaea. [6]

A new group of FBPases (FBPase V) is found in thermophilic archaea and the hyperthermophilic bacterium Aquifex aeolicus. [7] The characterised members of this group show strict substrate specificity for FBP and are suggested to be the true FBPase in these organisms. [7] [8] A structural study suggests that FBPase V has a novel fold for a sugar phosphatase, forming a four-layer alpha-beta-beta-alpha sandwich, unlike the more usual five-layered alpha-beta-alpha-beta-alpha arrangement. [8] The arrangement of the catalytic side chains and metal ligands was found to be consistent with the three-metal ion assisted catalysis mechanism proposed for other FBPases.

The fructose 1,6-bisphosphatases found within the Firmicutes (low GC Gram-positive bacteria) do not show any significant sequence similarity to the enzymes from other organisms. The Bacillus subtilis enzyme is inhibited by AMP, though this can be overcome by phosphoenolpyruvate, and is dependent on Mn(2+). [9] [10] Mutants lacking this enzyme are apparently still able to grow on gluconeogenic growth substrates such as malate and glycerol.

Click on genes, proteins and metabolites below to link to respective articles. [§ 1]

Fructose 1,6-bisphosphatase also plays a key role in hibernation, which requires strict regulation of metabolic processes to facilitate entry into hibernation, maintenance, arousal from hibernation, and adjustments to allow long-term dormancy. [11] [12] [13] During hibernation, an animal's metabolic rate may decrease to around 1/25 of its euthermic resting metabolic rate. [12] [13] [14] FBPase is modified in hibernating animals to be much more temperature sensitive than it is in euthermic animals. [11] [13] [14] FBPase in the liver of a hibernating bat showed a 75% decrease in Km for its substrate FBP at 5 °C than at 37 °C. [11] However, in a euthermic bat this decrease was only 25%, demonstrating the difference in temperature sensitivity between hibernating and euthermic bats. [11] When sensitivity to allosteric inhibitors such as AMP, ADP, inorganic phosphate, and fructose-2,6-bisphosphate were examined, FBPase from hibernating bats was much more sensitive to inhibitors at low temperature than in euthermic bats. [11] [15] [16]

During hibernation, respiration also dramatically decreases, resulting in conditions of relative anoxia in the tissues. Anoxic conditions inhibit gluconeogenesis, and therefore FBPase, while stimulating glycolysis, and this is another reason for reduced FBPase activity in hibernating animals. [17] The substrate of FBPase, fructose 1,6-bisphosphate, has also been shown to activate pyruvate kinase in glycolysis, linking increased glycolysis to decreased gluconeogenesis when FBPase activity is decreased during hibernation. [13]

In addition to hibernation, there is evidence that FBPase activity varies significantly between warm and cold seasons even for animals that do not hibernate. [18] In rabbits exposed to cold temperatures, FBPase activity decreased throughout the duration of cold exposure, increasing when temperatures became warmer again. [18] The mechanism of this FBPase inhibition is thought to be digestion of FBPase by lysosomal proteases, which are released at higher levels during colder periods. [18] Inhibition of FBPase through proteolytic digestion decreases gluconeogenesis relative to glycolysis during cold periods, similar to hibernation. [18]

Fructose 1,6-bisphosphate aldolase is another temperature dependent enzyme that plays an important role in the regulation of glycolysis and gluconeogenesis during hibernation. [14] Its main role is in glycolysis instead of gluconeogenesis, but its substrate is the same as FBPase's, so its activity affects that of FBPase in gluconeogenesis. Aldolase shows similar changes in activity to FBPase at colder temperatures, such as an upward shift in optimum pH at colder temperatures. This adaptation allows enzymes such as FBPase and fructose-1,6-bisphosphate aldolase to track intracellular pH changes in hibernating animals and match their activity ranges to these shifts. [14] Aldolase also complements the activity of FBPase in anoxic conditions (discussed above) by increasing glycolytic output while FBPase inhibition decreases gluconeogenesis activity. [19]

Fructose 1,6-bisphosphatase is also a key player in treating type 2 diabetes. In this disease, hyperglycemia causes many serious problems, and treatments often focus on lowering blood sugar levels. [20] [21] [22] Gluconeogenesis in the liver is a major cause of glucose overproduction in these patients, and so inhibition of gluconeogenesis is a reasonable way to treat type 2 diabetes. FBPase is a good enzyme to target in the gluconeogenesis pathway because it is rate-limiting and controls the incorporation of all three-carbon substrates into glucose but is not involved in glycogen breakdown and is removed from mitochondrial steps in the pathway. [20] [21] [22] This means that altering its activity can have a large effect on gluconeogenesis while reducing the risk of hypoglycemia and other potential side effects from altering other enzymes in gluconeogenesis. [20] [21]

Drug candidates have been developed that mimic the inhibitory activity of AMP on FBPase. [20] [22] Efforts were made to mimic the allosteric inhibitory effects of AMP while making the drug as structurally different from it as possible. [22] Second-generation FBPase inhibitors have now been developed and have had good results in clinical trials with non-human mammals and now humans. [20] [23]


Clark, S. L. Jr. Cellular differentiation in the kidneys of newborn mice studied with the electron microscope. J. Biophys. Biochem. Cytol. 3, 349–362 (1957).

Novikoff, A. B. The proximal tubule cell in experimental hydronephrosis. J. Biophys. Biochem. Cytol. 6, 136–138 (1959).

Ashford, T. P. & Porter, K. R. Cytoplasmic components in hepatic cell lysosomes. J. Cell Biol. 12, 198–202 (1962).

Novikoff, A. B. & Essner, E. Cytolysomes and mitochondrial degeneration. J. Cell Biol. 15, 140–146 (1962).

de Duve, C. & Wattiaux, R. Functions of lysosomes. Annu. Rev. Physiol. 28, 435–492 (1966).

Novikoff, A. B., Essner, E. & Quintana, N. Golgi apparatus and lysosomes. Fed. Proc. 23, 1010–1022 (1964).

Deter, R. L., Baudhuin, P. & de Duve, C. Participation of lysosomes in cellular autophagy induced in rat liver by glucagon. J. Cell Biol. 35, C11–16 (1967).

Pfeifer, U. Inhibition by insulin of the physiological autophagic breakdown of cell organelles. Acta Biol. Med. Ger. 36, 1691–1694 (1977).

Mortimore, G. E. & Schworer, C. M. Induction of autophagy by amino-acid deprivation in perfused rat liver. Nature 270, 174–176 (1977).

Seglen, P. O. & Gordon, P. B. 3-methyladenine: specific inhibitor of autophagic/lysosomal protein degradation in isolated rat hepatocytes. Proc. Natl Acad. Sci. USA 79, 1889–1892 (1982).

Holen, I., Gordon, P. B. & Seglen, P. O. Protein kinase-dependent effects of okadaic acid on hepatocytic autophagy and cytoskeletal integrity. Biochem. J. 284, 633–636 (1992).

Gordon, P. B. & Seglen, P. O. Prelysosomal convergence of autophagic and endocytic pathways. Biochem. Biophys. Res. Commun. 151, 40–47 (1988).

Bolender, R. P. & Weibel, E. R. A morphometric study of the removal of phenobarbital-induced membranes from hepatocytes after cessation of treatment. J. Cell Biol. 56, 746–761 (1973).

Beaulaton, J. & Lockshin, R. A. Ultrastructural study of the normal degeneration of the intersegmental muscles of Anthereae polyphemus and Manduca sexta (Insecta, Lepidoptera) with particular reference of cellular autophagy. J. Morphol. 154, 39–57 (1977).

Veenhuis, M., Douma, A., Harder, W. & Osumi, M. Degradation and turnover of peroxisomes in the yeast Hansenula polymorpha induced by selective inactivation of peroxisomal enzymes. Arch. Microbiol. 134, 193–203 (1983).

Lemasters, J. J. et al. The mitochondrial permeability transition in cell death: a common mechanism in necrosis, apoptosis and autophagy. Biochim. Biophys. Acta 1366, 177–196 (1998).

Takeshige, K., Baba, M., Tsuboi, S., Noda, T. & Ohsumi, Y. Autophagy in yeast demonstrated with proteinase-deficient mutants and conditions for its induction. J. Cell Biol. 119, 301–311 (1992).

Tsukada, M. & Ohsumi, Y. Isolation and characterization of autophagy-defective mutants of Saccharomyces cerevisiae. FEBS Lett. 333, 169–174 (1993).

Titorenko, V. I., Keizer, I., Harder, W. & Veenhuis, M. Isolation and characterization of mutants impaired in the selective degradation of peroxisomes in the yeast Hansenula polymorpha. J. Bacteriol. 177, 357–363 (1995).

Harding, T. M., Morano, K. A., Scott, S. V. & Klionsky, D. J. Isolation and characterization of yeast mutants in the cytoplasm to vacuole protein targeting pathway. J. Cell Biol. 131, 591–602 (1995).

Klionsky, D. J. et al. A unified nomenclature for yeast autophagy-related genes. Dev. Cell 5, 539–545 (2003).

Matsuura, A., Tsukada, M., Wada, Y. & Ohsumi, Y. Apg1p, a novel protein kinase required for the autophagic process in Saccharomyces cerevisiae. Gene 192, 245–250 (1997).

Kanki, T. et al. A genomic screen for yeast mutants defective in selective mitochondria autophagy. Mol. Biol. Cell 20, 4730–4738 (2009).

Okamoto, K., Kondo-Okamoto, N. & Ohsumi, Y. Mitochondria-anchored receptor Atg32 mediates degradation of mitochondria via selective autophagy. Dev. Cell 17, 87–97 (2009).

Klionsky, D. J., Cueva, R. & Yaver, D. S. Aminopeptidase I of Saccharomyces cerevisiae is localized to the vacuole independent of the secretory pathway. J. Cell Biol. 119, 287–299 (1992).

Hutchins, M. U. & Klionsky, D. J. Vacuolar localization of oligomeric α-mannosidase requires the cytoplasm to vacuole targeting and autophagy pathway components in Saccharomyces cerevisiae. J. Biol. Chem. 276, 20491–20498 (2001).

Xie, Z. & Klionsky, D. J. Autophagosome formation: core machinery and adaptations. Nat. Cell Biol. 9, 1102–1109 (2007).

Suzuki, K. et al. The pre-autophagosomal structure organized by concerted functions of APG genes is essential for autophagosome formation. EMBO J. 20, 5971–5981 (2001).

Kim, J., Huang, W. -P., Stromhaug, P. E. & Klionsky, D. J. Convergence of multiple autophagy and cytoplasm to vacuole targeting components to a perivacuolar membrane compartment prior to de novo vesicle formation. J. Biol. Chem. 277, 763–773 (2002).

Teter, S. A. et al. Degradation of lipid vesicles in the yeast vacuole requires function of Cvt17, a putative lipase. J. Biol. Chem. 276, 2083–2087 (2001).

Epple, U. D., Suriapranata, I., Eskelinen, E.-L. & Thumm, M. Aut5/Cvt17p, a putative lipase essential for disintegration of autophagic bodies inside the vacuole. J. Bacteriol. 183, 5942–5955 (2001).

Yang, Z., Huang, J., Geng, J., Nair, U. & Klionsky, D. J. Atg22 recycles amino acids to link the degradative and recycling functions of autophagy. Mol. Biol. Cell 17, 5094–5104 (2006).

Mizushima, N., Sugita, H., Yoshimori, T. & Ohsumi, Y. A new protein conjugation system in human. The counterpart of the yeast Apg12p conjugation system essential for autophagy. J. Biol. Chem. 273, 33889–33892 (1998).

Kabeya, Y. et al. LC3, a mammalian homologue of yeast Apg8p, is localized in autophagosome membranes after processing. EMBO J. 19, 5720–5728 (2000).

Klionsky, D. J. et al. Guidelines for the use and interpretation of assays for monitoring autophagy in higher eukaryotes. Autophagy 4, 151–175 (2008).

Yang, Z. & Klionsky, D. J. Mammalian autophagy: core molecular machinery and signaling regulation. Curr. Opin. Cell Biol. 22, 124–131 (2009).

Mizushima, N. et al. Dissection of autophagosome formation using Apg5-deficient mouse embryonic stem cells. J. Cell Biol. 152, 657–668 (2001).

Tian, Y. et al. C. elegans screen identifies autophagy genes specific to multicellular organisms. Cell 141, 1042–1055 (2010).

Behrends, C., Sowa, M. E., Gygi, S. P. & Harper, J. W. Network organization of the human autophagy system. Nature 466, 68–76 (2010).

Lipinski, M. M. et al. A genome-wide siRNA screen reveals multiple mTORC1 independent signaling pathways regulating autophagy under normal nutritional conditions. Dev. Cell 18, 1041–1052 (2010).

Axe, E. L. et al. Autophagosome formation from membrane compartments enriched in phosphatidylinositol 3-phosphate and dynamically connected to the endoplasmic reticulum. J. Cell Biol. 182, 685–701 (2008).

Hayashi-Nishino, M. et al. A subdomain of the endoplasmic reticulum forms a cradle for autophagosome formation. Nat. Cell Biol. 11, 1433–1437 (2009).

Yla-Anttila, P., Vihinen, H., Jokitalo, E. & Eskelinen, E.-L. 3D tomography reveals connections between the phagophore and endoplasmic reticulum. Autophagy 5, 1180–1185 (2009).

Hailey, D. W. et al. Mitochondria supply membranes for autophagosome biogenesis during starvation. Cell 141, 656–667 (2010).

Ravikumar, B., Moreau, K., Jahreiss, L., Puri, C. & Rubinsztein, D. C. Plasma membrane contributes to the formation of pre-autophagosomal structures. Nat. Cell Biol. 12, 747–757 (2010).

Tooze, S. A & Yoshimori, T. The origin of the autophagosomal membrane. Nat. Cell Biol. 12, 831–835 (2010).

Bavro, V. N. et al. Crystal structure of the GABAA-receptor-associated protein, GABARAP. EMBO Rep. 3, 183–189 (2002).

Sugawara, K. et al. The crystal structure of microtubule-associated protein light chain 3, a mammalian homologue of Saccharomyces cerevisiae Atg8. Genes Cells 9, 611–618 (2004).

Miller, S. et al. Shaping development of autophagy inhibitors with the structure of the lipid kinase Vps34. Science 327, 1638–1642 (2010).

Kunz, J. et al. Target of rapamycin in yeast, TOR2, is an essential phosphatidylinositol kinase homolog required for G1 progression. Cell 73, 585–596 (1993).

Brown, E. J. et al. A mammalian protein targeted by G1-arresting rapamycin-receptor complex. Nature 369, 756–758 (1994).

Blommaart, E. F., Luiken, J. J., Blommaart, P. J., van Woerkom, G. M. & Meijer, A. J. Phosphorylation of ribosomal protein S6 is inhibitory for autophagy in isolated rat hepatocytes. J. Biol. Chem. 270, 2320–2326 (1995).

Noda, T. & Ohsumi, Y. Tor, a phosphatidylinositol kinase homologue, controls autophagy in yeast. J Biol Chem 273, 3963–3966 (1998).

Blommaart, E. F., Krause, U., Schellens, J. P., Vreeling-Sindelarova, H. & Meijer, A. J. The phosphatidylinositol 3-kinase inhibitors wortmannin and LY294002 inhibit autophagy in isolated rat hepatocytes. Eur. J. Biochem. 243, 240–246 (1997).

Petiot, A., Ogier-Denis, E., Blommaart, E. F., Meijer, A. J. & Codogno, P. Distinct classes of phosphatidylinositol 3′-kinases are involved in signaling pathways that control macroautophagy in HT-29 cells. J. Biol. Chem. 275, 992–998 (2000).

Arico, S. et al. The tumor suppressor PTEN positively regulates macroautophagy by inhibiting the phosphatidylinositol 3-kinase/protein kinase B pathway. J. Biol. Chem. 276, 35243–35246 (2001).

Meijer, A. J. & Codogno, P. Regulation and role of autophagy in mammalian cells. Int. J. Biochem. Cell Biol. 36, 2445–2462 (2004).

Wei, Y., Pattingre, S., Sinha, S., Bassik, M. & Levine, B. JNK1-mediated phosphorylation of Bcl-2 regulates starvation-induced autophagy. Mol. Cell 30, 678–688 (2008).

Zalckvar, E. et al. DAP-kinase-mediated phosphorylation on the BH3 domain of beclin 1 promotes dissociation of beclin 1 from Bcl-XL and induction of autophagy. EMBO Rep. 10, 285–292 (2009).

Liang, X. H. et al. Induction of autophagy and inhibition of tumorigenesis by beclin 1. Nature 402, 672–676 (1999).

Pattingre, S. et al. Bcl-2 anti-apoptotic proteins inhibit Beclin 1-dependent autophagy. Cell 122, 927–939 (2005).

Aita, V. M. et al. Cloning and genomic organization of beclin 1, a candidate tumor suppressor gene on chromosome 17q21. Genomics 59, 59–65 (1999).

Qu, X. et al. Promotion of tumorigenesis by heterozygous disruption of the beclin 1 autophagy gene. J. Clin. Invest. 112, 1809–1820 (2003).

Yue, Z., Jin, S., Yang, C., Levine, A. J. & Heintz, N. Beclin 1, an autophagy gene essential for early embryonic development, is a haploinsufficient tumor suppressor. Proc. Natl Acad. Sci. USA 100, 15077–15082 (2003).

Marino, G. et al. Tissue-specific autophagy alterations and increased tumorigenesis in mice deficient in Atg4C/autophagin-3. J. Biol. Chem. 282, 18573–18583 (2007).

Degenhardt, K. et al. Autophagy promotes tumor cell survival and restricts necrosis, inflammation, and tumorigenesis. Cancer Cell 10, 51–64 (2006).

Mathew, R. et al. Autophagy suppresses tumor progression by limiting chromosomal instability. Genes Dev. 21, 1367–1381 (2007).

Mathew, R. et al. Autophagy suppresses tumorigenesis through elimination of p62. Cell 137, 1062–1075 (2009).

Amaravadi, R. K. et al. Autophagy inhibition enhances therapy-induced apoptosis in a Myc-induced model of lymphoma. J. Clin. Invest. 117, 326–336 (2007).

Carew, J. S. et al. Targeting autophagy augments the anticancer activity of the histone deacetylase inhibitor SAHA to overcome Bcr–Abl-mediated drug resistance. Blood 110, 313–322 (2007).

Ravikumar, B., Duden, R. & Rubinsztein, D. C. Aggregate-prone proteins with polyglutamine and polyalanine expansions are degraded by autophagy. Hum. Mol. Genet. 11, 1107–1117 (2002).

Ravikumar, B. et al. Inhibition of mTOR induces autophagy and reduces toxicity of polyglutamine expansions in fly and mouse models of Huntington disease. Nat. Genet. 36, 585–595 (2004).

Webb, J. L., Ravikumar, B., Atkins, J., Skepper, J. N. & Rubinsztein, D. C. α-Synuclein is degraded by both autophagy and the proteasome. J. Biol. Chem. 278, 25009–25013 (2003).

Yu, W. H. et al. Macroautophagy—a novel β-amyloid peptide-generating pathway activated in Alzheimer's disease. J. Cell Biol. 171, 87–98 (2005).

Komatsu, M. et al. Loss of autophagy in the central nervous system causes neurodegeneration in mice. Nature 441, 880–884 (2006).

Hara, T. et al. Suppression of basal autophagy in neural cells causes neurodegenerative disease in mice. Nature 441, 885–889 (2006).

Bjørkøy, G. et al. p62/SQSTM1 forms protein aggregates degraded by autophagy and has a protective effect on huntingtin-induced cell death. J. Cell. Biol. 171, 603–614 (2005).

Iwata, A., Riley, B. E., Johnston, J. A. & Kopito, R. R. HDAC6 and microtubules are required for autophagic degradation of aggregated huntingtin. J. Biol. Chem. 280, 40282–40292 (2005).

Komatsu, M. et al. Homeostatic levels of p62 control cytoplasmic inclusion body formation in autophagy-deficient mice. Cell 131, 1149–1163 (2007).

Kirkin, V. et al. A role for NBR1 in autophagosomal degradation of ubiquitinated substrates. Mol. Cell 33, 505–516 (2009).

Nezis, I. P. et al. Ref(2)P, the Drosophila melanogaster homologue of mammalian p62, is required for the formation of protein aggregates in adult brain. J. Cell Biol. 180, 1065–1071 (2008).

Pankiv, S. et al. p62/SQSTM1 binds directly to Atg8/LC3 to facilitate degradation of ubiquitinated protein aggregates by autophagy. J. Biol. Chem. 282, 24131–24145 (2007).

Kim, P. K., Hailey, D. W., Mullen, R. T. & Lippincott-Schwartz, J. Ubiquitin signals autophagic degradation of cytosolic proteins and peroxisomes. Proc. Natl Acad. Sci. USA 105, 20567–20574 (2008).

Kraft, C., Peter, M. & Hoffman, K. Selective autophagy: ubiquitin-mediated recognition and beyond. Nat. Cell Biol. 12, 836–841 (2010).

Rikihisa, Y. Glycogen autophagosomes in polymorphonuclear leukocytes induced by rickettsiae. Anat. Rec. 208, 319–327 (1984).

Gutierrez, M. G. et al. Autophagy is a defense mechanism inhibiting BCG and Mycobacterium tuberculosis survival in infected macrophages. Cell 119, 753–766 (2004).

Nakagawa, I. et al. Autophagy defends cells against invading group A Streptococcus. Science 306, 1037–1040 (2004).

Andrade, R. M., Wessendarp, M., Gubbels, M. J., Striepen, B. & Subauste, C. S. CD40 induces macrophage anti-Toxoplasma gondii activity by triggering autophagy-dependent fusion of pathogen-containing vacuoles and lysosomes. J. Clin. Invest. 116, 2366–2377 (2006).

Singh, S. B., Davis, A. S., Taylor, G. A. & Deretic, V. Human IRGM induces autophagy to eliminate intracellular mycobacteria. Science 313, 1438–1441 (2006).

Ogawa, M. et al. Escape of intracellular Shigella from autophagy. Science 307, 727–731 (2005).

Yano, T. et al. Autophagic control of Listeria through intracellular innate immune recognition in Drosophila. Nat. Immunol. 9, 908–916 (2008).

Thurston, T. L., Ryzhakov, G., Bloor, S., von Muhlinen, N. & Randow, F. The TBK1 adaptor and autophagy receptor NDP52 restricts the proliferation of ubiquitin-coated bacteria. Nat. Immunol. 10, 1215–1221 (2009).

Liang, X. H. et al. Protection against fatal Sindbis virus encephalitis by beclin, a novel Bcl-2-interacting protein. J. Virol. 72, 8586–8596 (1998).

Tallóczy, Z., Virgin, H. W. IV. & Levine, B. PKR-dependent autophagic degradation of herpes simplex virus type 1. Autophagy 2, 24–29 (2006).

Orvedahl, A. et al. HSV-1 ICP34.5 confers neurovirulence by targeting the Beclin 1 autophagy protein. Cell Host Microbe 1, 23–35 (2007).

Paludan, C. et al. Endogenous MHC class II processing of a viral nuclear antigen after autophagy. Science 307, 593–596 (2005).

English, L. et al. Autophagy enhances the presentation of endogenous viral antigens on MHC class I molecules during HSV-1 infection. Nat. Immunol. 10, 480–487 (2009).

Del Roso, A. et al. Ageing-related changes in the in vivo function of rat liver macroautophagy and proteolysis. Exp. Gerontol. 38, 519–527 (2003).

Donati, A. et al. Age-related changes in the regulation of autophagic proteolysis in rat isolated hepatocytes. J. Gerontol. A Biol. Sci..Med. Sci. 56, B288–293 (2001).

Cavallini, G., Donati, A., Gori, Z. & Bergamini, E. Towards an understanding of the anti-aging mechanism of caloric restriction. Curr. Aging Sci. 1, 4–9 (2008).

Meléndez, A. et al. Autophagy genes are essential for dauer development and life-span extension in C. elegans. Science 301, 1387–1391 (2003).

Juhasz, G., Erdi, B., Sass, M. & Neufeld, T. P. Atg7-dependent autophagy promotes neuronal health, stress tolerance and longevity but is dispensable for metamorphosis in Drosophila. Genes Dev. 21, 3061–3066 (2007).

Simonsen, A. et al. Promoting basal levels of autophagy in the nervous system enhances longevity and oxidant resistance in adult Drosophila. Autophagy 4, 176–184 (2008).

Madeo, F., Tavernarakis, N. & Kroemer, G. Can autophagy promote longevity? Nat. Cell Biol. 12, 842–846 (2010).

Bergamini, E. Targets for antiageing drugs. Expert Opin. Ther. Targets 9, 77–82 (2005).

Eisenberg, T. et al. Induction of autophagy by spermidine promotes longevity. Nat. Cell Biol. 11, 1305–1314 (2009).

Otto, G. P., Wu, M. Y., Kazgan, N., Anderson, O. R. & Kessin, R. H. Macroautophagy is required for multicellular development of the social amoeba Dictyostelium discoideum. J. Biol. Chem. 278, 17636–17645 (2003).

Juhasz, G., Csikos, G., Sinka, R., Erdelyi, M. & Sass, M. The Drosophila homolog of Aut1 is essential for autophagy and development. FEBS Lett. 543, 154–158 (2003).

Kuma, A. et al. The role of autophagy during the early neonatal starvation period. Nature 432, 1032–1036 (2004).

Komatsu, M. et al. Impairment of starvation-induced and constitutive autophagy in Atg7-deficient mice. J. Cell Biol. 169, 425–434 (2005).

Tsukamoto, S. et al. Autophagy is essential for pre-implantation development of mouse embryos. Science 321, 117–120 (2008).

Mizushima, N. & Levine, B. Autophagy in mammalian development and differentiation. Nat. Cell Biol. 12, 823–830 (2010).

Schin, K. S. & Clever, U. Lysosomal and free acid phosphatase in salivary glands of Chironomus tentans. Science 150, 1053–1055 (1965).

Nopanitaya, W. & Misch, D. W. Developmental cytology of the midgut in the flesh-fly, Sarcophaga bullata (Parker). Tissue Cell 6, 487–502 (1974).

Yu, L. et al. Regulation of an ATG7–beclin 1 program of autophagic cell death by caspase-8. Science 304, 1500–1502 (2004).

Shimizu, S. et al. Role of Bcl-2 family proteins in a non-apoptotic programmed cell death dependent on autophagy genes. Nat. Cell Biol. 6, 1221–1228 (2004).

Berry, D. L. & Baehrecke, E. H. Growth arrest and autophagy are required for salivary gland cell degradation in Drosophila. Cell 131, 1137–1148 (2007).

McPhee, C. K., Logan, M. A., Freeman, M. R. & Baehrecke, E. H. Activation of autophagy during cell death requires the engulfment receptor Draper. Nature 465, 1093–1096 (2010).

Mathew, R., Karantza-Wadsworth, V. & White, E. Role of autophagy in cancer. Nat. Rev. Cancer 7, 961–967 (2007).

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