Metabolomic profiling reveals developmentally regulated biosynthesis of polyprenols and dolichols in the malaria parasite

The cis-polyisoprenoid lipids namely polyprenols (POH), dolichols (DOH) and their derivatives are linear polymers of several isoprene units and have recently gained special attention due to several breakthroughs in the field including improvements in the analytical techniques for their analysis, partial identification of their enzymatic machinery, and the discovery of new biological functions beyond glycosylation. In eukaryotes, POH and DOH are synthesized as a mixture of four or more homologues of different length with one or two predominant species with sizes varying among organisms. Interestingly, POH are hardly detectable in eukaryotic cells under normal conditions with the exception of plants and sporulating yeast. Our metabolomics studies revealed that cis-polyisoprenoids are more prevalent and diverse in the protozoan parasite Plasmodium falciparum than previously postulated as we uncovered the active biosynthesis of medium-long POH and DOH (15 to 19 isoprene units). Moreover, a distinctive POH and DOH profile both within the intraerythrocytic asexual cycle and between asexual and gametocyte stages was observed. These results suggest that cis-polyisoprenoid biosynthesis is developmentally regulated. In addition, we confirmed that DOH biosynthesis occurs via reduction of the POH to DOH by an active polyprenol reductase (PfPPRD) and metabolomics analyses of a PfPPRD conditional mutant suggest that a salvage mechanism of DOH may exist.

polyisoprenoid biosynthesis is developmentally regulated. In addition, we confirmed that DOH biosynthesis occurs via reduction of the POH to DOH by an active polyprenol reductase (PfPPRD) and metabolomics analyses of a PfPPRD conditional mutant suggest that a salvage mechanism of DOH may exist.
Malaria is caused by protozoan parasites of the genus Plasmodium and most cases of lifethreatening malaria are attributable to infection with Plasmodium falciparum. The parasite has a complex life cycle that involves the human host and its vector, the Anopheles mosquitoes. All clinical features are caused during the asexual intraerythrocytic life cycle due to the repeated invasion of human red blood cells (RBCs). The intraerythrocytic cycle of P. falciparum lasts around 48 h, during which the parasite progresses through four morphologically different stages: ring, trophozoite, and schizont stages, ending with the rupture of the erythrocyte and release of merozoites that will invade new erythrocytes. Transmission of the malaria parasite requires the development of male and female gametocytes (gametocytogenesis), which are ingested by female mosquitoes during a blood meal and undergo sexual reproduction in the mosquito's midgut. Nondividing P. falciparum gametocytes take between 10 and 12 days to fully mature and progress through five morphologically distinct forms (stages I to V), which are different from other Plasmodium species.
Over one billion people are at risk of contracting malaria. In 2016, an increase of 5 million malaria cases were reported, mainly in the Americas, and 98 million additional treatment courses of artemisinin-based combination therapy (ACT) were obtained by countries as compared to 2015 (1). ACT combines two or more drugs with different modes of action with artemisinin and is the first-line treatment of P. falciparum infection. Unfortunately, the rapid development of resistance to both the partner drug and now to artemisinin supports the urgent need for identifying different therapeutic targets to expand our repertoire of antimalarials (2).
Historically, metabolic pathways have been a source of therapeutic targets especially for infectious diseases when pathways or their enzymes are sufficiently different or absent in the human host. The mature RBCs are capable of only a few metabolic functions since they do not contain organelles. Among the metabolic pathways that are not active in mature RBCs is the mevalonate pathway which synthesizes the isoprenoid building blocks isopentenyl diphosphate (IPP) and dimethylallyl diphosphate (DMAPP). The downstream cis-polyisoprenoid biosynthesis for dolichols that occurs mainly in the endoplasmic reticulum (ER) is also inactive in mature RBCs (3). On the other hand, P. falciparum has active isoprenoid biosynthesis during the asexual intraerythrocytic cycle as well as during gametocytogenesis where the isoprenoid precursors are synthesized through the methylerythritol phosphate (MEP) pathway (4)(5)(6).
The MEP pathway is localized in the apicoplast (7,8), a unique chloroplast-like organelle essential for growth and pathogenesis of the malaria parasite (9) (Fig.1). Moreover, supply of the isoprenoid precursor IPP is the sole metabolic function of the apicoplast in asexual intraerythrocytic and gametocyte stages (6,10). The fact that IPP alone allows parasites lacking the apicoplast to normally grow and develop indicates that IPP is transported out of the apicoplast where the synthesis of cis-polyisoprenoid products is predicted to occur (Fig. 1). cis-Polyisoprenoids (polyprenols (POH), dolichols (DOH) and their derivatives) are linear polymers of several isoprene units. These lipids are present in all membrane systems (11,12) but their biological functions besides protein glycosylation in the ER remains largely unknown. Recently, cis-polyisoprenoids gained special attention due to several breakthroughs in the field including improvements in the analytical techniques for their analysis (13), partial identification of their enzymatic machinery (14), and the discovery of new biological functions beyond glycosylation (15,16). Among some of these biological functions are the regulation of the membrane fluidity (16), stimulation of spore wall formation in yeast (15), and scavenging free radicals in cell membranes (17)(18)(19). However, cispolyisoprenoid metabolism and its biological functions in P. falciparum remain poorly understood.
The initial step of cis-polyisoprenoid biosynthesis (POH and DOH) in the malaria parasite is common to all isoprenoid products (Fig.  1) where a multifunctional enzyme synthesizes farnesyl diphosphate (FPP) and geranylgeranyl diphosphate (GGPP) (20). Synthesis of cispolyprenyl-diphosphate (PDP) is then catalyzed by a cis-prenyltransferase (CPT) which adds several IPP molecules in cis-configuration to FPP or GGPP. This step is called elongation and in eukaryotes occurs in the ER where their downstream products dolichyl phosphates (Dol-P) serve as lipid linked oligosaccharide (LLO) carrier for protein glycosylation and glycosylphosphatidylinositol (GPI) synthesis. In other organisms, cis-polyisoprenoid synthesis also occurs in peroxisomes (rat (21), yeast (22)), lipid droplets (yeast (15,23)) and chloroplasts (plants, cyanobacteria (16,24)). The postulated final steps of cis-polyisoprenoid biosynthesis involve dephosphorylation of PDP to POH followed by reduction of the a-isoprene unit by polyprenol reductase (PPRD), also called SRD5A3 or DFG10 in mammals and yeast, respectively. Whether dephosphorylation of PDP is catalyzed by one or two enzymes (25) has not been elucidated in any organism. Moreover, it has also been suggested that an isopentenol or isopentanol, instead of IPP, is added at the last step, which would avoid the need of dephosphorylation or dephosphorylation and reduction, respectively, but it remains to be addressed (11,26).
In eukaryotes, DOH and POH are synthesized as a mixture of four or more different lengths (indicated by the total number of carbons or IPP units, e.g. POH/DOH 11 or C55) with one or two predominant species where the size varies among organisms. Interestingly, POH are hardly detectable in eukaryotic cells under normal conditions with the exception of plant roots and sporulating yeast. Previously, it was reported that the malaria parasite synthesizes DOH of 11 and 12 isoprene units (27) and their biosynthesis peaks at the schizont stage (4,5). Here, our metabolomic studies uncovered active biosynthesis of mediumlong POH and DOH (15 to 19 isoprene units) in the malaria parasite as well as a distinctive POH and DOH profile within the intraerythrocytic asexual cycle and between asexual and gametocyte stages. In addition, we confirmed that DOH biosynthesis occurs via reduction of the POH to DOH by an active PfPPRD (PF3D7_1455900) and metabolic analyses in a conditional knockdown of PfPPRD suggest the presence of a salvage mechanism of DOH.

Results and Discussion
Medium-long polyprenols and dolichols are present in the asexual and sexual intraerythrocytic stages of P. falciparum Dolichol biosynthesis in the malaria parasites has been assessed previously using radiolabeled metabolic precursors followed by TLC or HPLC analysis. Thus, in order to expand our knowledge of cis-polyisoprenoids present in the malaria parasite especially in gametocyte stages, we standardized an untargeted metabolomics approach using ion-key separation, a high-pressure microfluidic device that provides sub-2-µm Ultra-Performance Liquid Chromatography (UPLC) separations with the SYNAPT G2-Si mass spectrometer, enabling highly sensitive and efficient microflow LC-HRMS (High Resolution Mass Spectrometry) separations that enhance detection of metabolites (see experimental procedures). Surprisingly, we identified a distinctive POH and DOH profile between asexual and gametocyte stages that also differs from the human erythrocyte cell host (Fig. 2). To our knowledge, this is the first time that detectable levels of POH are reported to be present in another eukaryotic cell besides plants and sporulating yeast.
Our studies revealed that cispolyisoprenoids are more prevalent and diverse in the malaria parasite than previously postulated (27,28). In the asexual intraerythrocytic stages, we detected the presence of both POH and DOH of 15 to 19 isoprene units, with 15-, 16-and 17-isoprene units being the predominant species (Fig. 2). Interestingly, ring and trophozoite stages presented higher DOH/POH ratios than schizonts (Table 1). In stage IV gametocytes, DOH of 15 to 20 isoprene units were also present, but the predominant species are 17-, 18-, 19-isoprene units (Fig. 2). Interestingly, POH were difficult to detect in gametocytes under our experimental conditions. As expected, human RBCs contain low levels of only DOH of 18 to 20 isoprene units. This is the first time that the cis-polyisoprenoid profile is determined by LC-HRMS in P. falciparum, and under our experimental conditions DOH and POH of 11 and 12 were not detected probably due to their low abundance.
The cis-polyisoprenoid lipids are present in all membrane systems (11,12) but their biological functions besides protein glycosylation in the ER remain largely unknown. A recent fluorescence anisotropy analysis performed in the protein-dense thylakoid membranes of the chloroplast, revealed that membrane areas with less POH had greater membrane fluidity and photosynthesis was affected (16). These analyses suggest that POH have an effect on the biophysical characteristics of biological membranes. Moreover, it was recently demonstrated in yeast that lipid droplets of vegetative cells contain mainly DOH of 14-17 isoprene units while sporulating cells not only shift toward longer species (19 to 24 isoprene units) but also accumulate similar levels of long POH (19 to 24 isoprene units). Long-chain POH synthetized by a developmentally regulated CPT present in lipid droplets (Srt1) promote spore wall formation by serving as a signal that activates chitin synthase (directly or indirectly which remains to be determined) possibly through changes in the membrane structure introduced by POH (15). Therefore, the presence of a distinctive POH and DOH profile within the intraerythrocytic asexual cycle and between asexual and gametocyte stages suggests that cis-polyisoprenoid biosynthesis is developmentally regulated and that these lipids may be involved in other cellular processes besides glycosylation.

Medium-long polyprenols and dolichols are synthesized de novo by P. falciparum
In order to confirm that the medium-long POH and DOH detected were in fact synthesized by the parasite, de novo biosynthesis was assessed by metabolic labeling using [1-13 C]glucose and [3-13 C]IPP (Fig. 3A). Glucose was selected as metabolic precursor for cis-polyisoprenoid biosynthesis because the mevalonate and MEP pathway-specific labeling patterns of isoprene units resulting from glucose metabolism via glycolysis are well established (13,29). In addition, the mevalonate pathway is inactive in the host cell, the human RBCs, thus, only MEP pathway-specific labeling patterns are observed.
Highly synchronous infected RBC in ring stage were first incubated in glucose-free media for 2 h and then supplemented with [1-13 C]glucose (4 g/L) for 24 h which is metabolized into [1,5-13 C]IPP (Fig. 3A). A similar scheme was used for the metabolic precursor [3-13 C]IPP, but in this case cultures were treated for 2 h with 10 µM fosmidomycin (FOS), an inhibitor of 1-deoxy-Dxylulose 5-phosphate reductoisomerase in the MEP pathway (DOXP ® MEP) (4,8), to reduce >75% the endogenous IPP pool as we reported previously (30), and then supplemented with [3-13 C]IPP: 12 C5-IPP (1:1, 200 µM final concentration) in the presence of FOS. As previously described (13,29), we observed 13 C-enrichment as a Gaussian distribution for both 13 C-precursors ( Fig. 3B and S1), where m/z shift is determined by the remaining of unlabeled metabolic precursor pools in the cell and the number of 13 C-atoms incorporated into IPP based on the isotopic dilution due to metabolism (13,31). As expected, only unlabeled DOH was detected in uninfected RBCs since there is no active de novo biosynthesis of isoprenoids (Fig. 3C). Thus, these results confirmed that the detected POH and DOH of 15 to 19 isoprene units are biosynthesized by the malaria parasite.
During optimization of the 13 C-biolabeling experimental design, we established that in order to observe 13 C-enrichment in POH and DOH, parasites required at least 20-24 h of growth in the presence of the 13 C-precursor starting at ring stage. Interestingly, independently of the metabolic 13 Cprecursor used, unlabeled DOH was always detected while unlabeled POH was indistinguishable from noise in the presence of a 13 C-precursor (Fig. 3B). In addition, we showed that POH and DOH achieved similar levels as the parasite progress from ring to the schizont stage. Altogether, these results suggest that synthesis of POH and DOH is developmentally regulated and that a reserve of cis-polyisoprenoids may be carried over to the next asexual cycle. It has been demonstrated that stage III gametocytes incorporate and metabolize 13 Cglucose (32). In addition, we previously reported that the MEP pathway intermediates are present in late-stage gametocytes and that IPP synthesis is essential for normal gametocytogenesis since gametocytes lacking the apicoplast require exogenous IPP supplementation until they reach stage IV (6). Surprisingly, 13 C-enrichment in DOH was not detected in stage IV gametocytes when [1-13 C]glucose was supplemented starting at stage III (Fig. 4). Therefore, these results suggest that synthesis of DOH occurs earlier than stage III in the gametocytogenesis or potentially these lipids are carried over from the previous asexual cycle and that IPP is required for other isoprenoid products than DOH in gametocyte stages III to IV (6).

P. falciparum polyprenol reductase restored glycosylation of mCPY and reduced POH levels in yeast dfg10D D mutant cells.
We hypothesized that the presence of POH in P. falciparum indicates that DOH biosynthesis may occur via reduction of the POH to DOH by an active PfPPRD (PF3D7_1455900) (Fig. 1). Therefore, the coding sequence of the putative PfPPRD was optimized for codon usage in yeast and its enzymatic function was confirmed by complementation assays in Saccharomyces cerevisiae. It was previously reported by Cantagrel and colleagues (33) that human PPRD (SRD5A3) can complement the N-glycosylation defects caused by an inactive yeast DFG10, which is the PPRD in this organism for DOH synthesis. As mentioned above, their downstream products Dol-P serve as lipid linked oligosaccharide carrier for protein glycosylation (Fig. 1). In this work, we used the dfg10D mutant cells where the DFG10 gene is deleted from the chromosome and displays a lack of N-glycosylation. Mature carboxypeptidase Y (mCPY) has four N-glycosylation sites which are all occupied under normal growth conditions in yeast (34) and depend on a functional PPRD (Fig.  5A) (33). A single band is observed in wild-type (WT) cells when mCPY is fully glycosylated while four extra bands were detected in dfg10D mutant cells representing the different glycoforms (-1, -2, -3, -4) (Fig. 5A, lane 2). The mutant transformed with yeast DGF10 or PfPPRD under the control of the endogenous DFG10 promoter showed full correction of the CPY underglycosylation. Some underglycosylated CPY was still detected in cells expressing PfPPRD suggesting that this enzyme was not as active as yeast DFG10.
In order to confirm the biochemical effects underlying the yeast complementation, we assessed the POH and DOH profile by LC-HRMS (Fig. 5B). As described previously (3,33), only DOH 15 and 16 were detected in wildtype (WT) yeast while POH along with DOH were detected in the dfg10D mutant cells. As expected, dfg10D mutant cells expressing PfPPRD showed reduction of the POH levels confirming its predicted enzymatic function (POH ® DOH).

Changes in the POH and DOH profiles without major growth defects suggests the presence of a salvage pathway
Two recent functional profiling studies of the Plasmodium genome indicate that Plasmodium PPRD is predicted to be essential (35,36). Thus, we used the CRISPR/Cas9 and the TetR-DOZI conditional system (37) to generate a transgenic strain for inducible knockdown of PfPPRD to assess if changes in the DOH/POH ratios may affect the in vitro growth of P. falciparum (Fig. 6A). Synthesis of DOH is predicted to occur in the ER, thus, a triple-HA tag was added to the C-terminus of PfPPRD to assess its localization. However, multiple attempts to tag PfPPRD with the TetR-DOZI-HA construct failed (Fig. S2), suggesting that PfPPRD tagged at the C-terminus is not efficiently translated or the protein is unstable. It is also possible that the highly conserved catalytic domain of PfPPRD is located in the C-terminal as predicted for the human and plant orthologs (33,38) and tagging interferes with proper enzymatic function. Therefore, we designed a similar construct (PfPPRD-TetR-DOZI) lacking a Cterminal tag. Using the untagged construct, we successfully modified the PfPPRD genomic locus and confirmed correct integration by PCR (Fig.  6B). The conditional knockdown system was induced by removing anhydrotetracycline (aTc), a stabilizer of the TetR-DOZI complex (37). The absence of aTc allows the TetR-DOZI complex to bind at the repetitive aptamer region reducing the translation of PfPPRD enzyme (Fig. 7A). Interestingly, a slight but significant growth defect was observed only after 8 days (5 th intraerythrocytic life cycle) without aTc. At this point certain morphological defects started to be observed in parasites (Fig. 7B). In the absence of PfPPRD tagging, it was not possible to determine whether the observed growth and morphological defects were attributed to a gradual or incomplete reduction in PfPPRD protein levels. Instead, the levels of POH and DOH were determined by LC-HRMS to assess whether PfPPRD enzymatic activity was reduced in the absence of aTc confirming that PfPPRD synthesizes DOH in vivo. These analyses were performed in trophozoite stage parasites where the highest DOH/POH ratios are observed (Table 1). Parasites grown in the absence of aTc for 3 days displayed a pronounced accumulation of POH 15 to 18 and a slight reduction of DOH 15 to 17 without changes in the levels of DOH 18 to 20 suggesting that some residual PfPPRD activity may still be present (Fig. 7C). Surprisingly, LC-HRMS analyses performed after at least 10 days in the absence of aTc revealed similar pronounced accumulation of POH 15 to 18 but DOH 15, 16 and 17 were reduced to undetectable levels while DOH 18, 19 and 20 were not significantly reduced under our experimental conditions (Fig. 7D). The observed reduction of DOH 15, 16 and 17 after 3 days of aTc removal was not statistically significant, but these lipids were undetectable after 10 days in the absence of aTc. Metabolic labeling experiments using [1-13 C]glucose in cultures after 10 days of aTc removal revealed that while 13 Cenrichment was observed in POH 18 supporting that PfCPT is active, 13 C-enrichment in DOH 18 was reduced to undetectable levels. Therefore, the detected DOH 18 was not biosynthesized de novo by parasites under our experimental conditions (Fig. S3). Moreover, 13 C-enrichment in DOH 19 was not detected in cultures where aTc was not removed. This result was not unexpected since low levels of 13 C-enrichment using [1-13 C]glucose were also observed using wild type parasites (Fig. S1). Since DOH 18, 19 and 20 are the species found in uninfected RBCs (Fig. 2), it is possible that the malaria parasite is able to uptake or sequester DOH species from the erythrocyte host but it becomes detectable, or probably intensified, when de novo DOH biosynthesis is impaired such as is the case of reduced PfPPRD activity. However, we cannot rule out that the parasite accumulated or salvaged these DOH species from its own biosynthesis as a result of reduced PfPPRD activity overtime which was negligible when 13 C-biolabeling was performed.
Since the biological role of these lipids remains largely unknown, it cannot be ruled out that POH may be able to perform some of the functions of DOH when their synthesis is compromised. Moreover, as mentioned above, areas of thylakoid membranes of a chloroplast with less POH had greater membrane fluidity and photosynthesis was affected (16). Therefore, it is also possible that the higher levels of POH detected in the absence of aTc may had affected membranes and their functions, thus, contributing to the growth and morphological defects observed.

Conclusion
Our studies using 13 C-profiling by LC-HRMS uncovered for the first time in P. falciparum co-occurrence of POH and DOH as well as synthesis of longer DOH species than previously reported. The presence of a distinctive POH and DOH profile within the intraerythrocytic asexual cycle and between asexual and gametocyte stages suggest that cis-polyisoprenoid biosynthesis is developmentally regulated. Interestingly, in mammals and plants increased level of POH results in aberrant glycosylation (39). However, it is difficult to assess if a similar response occurs in the malaria parasite since N-glycosylation in Plasmodium has been controversial despite the presence of very short N-glycans was previously reported in this parasite (40,41). Nevertheless, it is possible that the co-occurrence of POH and DOH suggest that these lipids may be involved in other cellular processes beside N-glycosylation as has been recently demonstrated in yeast (15) and plants (16) which guarantees further investigation.
The presence of POH in P. falciparum supports that DOH biosynthesis occurs via reduction of the POH to DOH and this step is catalyzed by PfPPRD (PF3D7_1455900). Interestingly, a knockdown of PfPPRD revealed that after aTc has been removed for several days, shorter DOH were undetectable while unlabeled DOH of 18 to 19 isoprene units were still present at which point a slight but significant growth defect as well as the observation of abnormal parasites were observed. Altogether, these results suggest that a salvage mechanism for DOH may be present and further studies are needed to establish their source (endogenous vs the host cell). Interestingly, P. falciparum cannot synthesize cholesterol (42), another isoprenoid product. Instead, the malaria parasite salvages cholesterol from its mammalian host (43,44) supporting the hypothesis that DOH salvage from RBCs may also occur.
As a result of several breakthroughs in the field of cis-polyisoprenoids (13)(14)(15), new biological functions of these lipids besides their known role in protein glycosylation are starting to be unveiled. Further analyses are still needed to answer why detectable levels of POH are present in the malaria parasite as well as why cis-polyisoprenoid species change during the parasite's life cycle which may reveal new biological functions of these unique lipids.

Stable isotope labeling and metabolite extraction of P. falciparum-infected and uninfected RBCs for polyisoprenoids profiling
Mycoplasma-free parasite cultures were tightly synchronized by 5% sorbitol treatment. Each biological replicate was obtained from 75 mL of P. falciparum-infected RBC cultures at 5% hematocrit and 7% parasitemia. For stage-specific polyisoprenoid profiling, parasites in trophozoite and schizont stages were obtained from P. falciparum-infected RBC cultures starting with early ring stage at 7% parasitemia and harvesting parasites at each indicated stage. Parasites at ring stage were obtained from synchronized cultures starting with early rings of P. falciparum at 3% parasitemia and recovering parasites after 48 h incubation to avoid collecting cells after sorbitol treatment.
For metabolic labeling experiments using [1-13 C]glucose, cultures in ring stage (5% hematocrit and 7% parasitemia) were first incubated in glucose-free medium for 2 h and then supplemented with 4 g/L of [1- 13  In order to assess the DOH/POH profile in early trophozoite culture of the PfPPRD conditional knockdown, parasites were grown in media supplemented either with 2.5 µg/mL Blasticidin S and 0.5 µM aTc, or only 2.5 µg/mL Blasticidin S for the indicated times (Fig. 7). 13 C-Biolabeling experiments were performed under similar conditions to induce the knockdown and then metabolically labeled as described above.
In all cases, infected RBCs were released from the host cell by lysis with 0.03% saponin in cold phosphate-buffered saline (PBS) solution containing 2 g/L glucose and washed three times with PBS/glucose by centrifuging 7 min at 10,000 × g at 4 o C and 10 µL were collected in the last wash to count parasites using a Countess cell counter (Thermo Fisher). Cell pellets were flash-frozen and stored at -80°C until metabolite extractions were performed.
Gametocyte stages were obtained using P. falciparum NF54 strain. Briefly, highly synchronous cultures at early ring stage were obtained by sorbitol treatment and purified using a MACS Ò magnetic affinity column (day 0) and set at 8-12% parasitemia and 5% hematocrit. On day 0 and 1, gametocytogenesis was induced by incubating cultures with 50% spent media and 50% fresh media for 40-48 h, as described previously (45). After reinvasion, asexual parasites were depleted from culture by addition of fresh media containing 20 U/mL of heparin for 4 days (day 2 to 5) and sorbitol treatment (46). After day 5, the media was replaced every two days until gametocytes reached stage IV. Development of parasites was monitored by microscopic evaluation of Giemsa-stained thin smears. For metabolic labeling experiments, stage III gametocyte cultures where obtained as described above (day 6 after induction). Then, parasites were incubated in glucose-free media for 2 h before adding [1-13 C]glucose (4 g/L) and incubated for 24 h when media was changed to add fresh [1-13 C]glucose (4 g/L) and to complete a total of 48 h labeling until gametocytes reached stage IV.
Extraction of metabolites was performed by treating samples with 1 mL cold methanol and three consecutive extractions with 2 mL of hexane followed by pulse-vortex for 1 min and 10 min sonication in an ultrasonic bath. The upper phases were combined, dried under nitrogen and stored at -80°C until LC-HRMS analysis. For LC-HRMS analysis, samples were resuspended in 100 µL of methanol/acetonitrile/2-propanol (60:25:15, v/v) pulse vortex 5 times and sonicate for 10 min.

Ultra-performance liquid chromatography and high-resolution mass spectrometry analysis
Analysis of polyisoprenoids was performed on an ionKey/MS system composed of an ACQUITY UPLC M-Class, the ionKey source, and an iKey HSS T3, 100 Å, 1.8 µm (particle size), 150 µm × 100 mm column coupled to a SYNAPT G2-Si mass spectrometer (Waters Corporation, Milford, MA, USA). Isoprenoid separation was accomplished using a binary gradient system consisting of methanol/acetonitrile (3:1, v/v), with 10 mM ammonium formate + 0.1% formic acid (mobile phase A) and 2-propanol/acetonitrile (3:1, v/v) containing 10 mM ammonium formate + 0.1% formic acid (mobile phase B). Sample analysis was performed using a linear gradient over a total run time of 35 min. The gradient was programmed as follows: 0.0-2.0 min 18% B, 2-16 min from 18% B to 89% B, 16-20 min at 89 % B, 20-25 min from 89% B to 18% B, and re-equilibrated for 10 min. The flow rate was 3 µL/min, the injection volume was 5 µL in full loop mode, and the ionKey temperature was set at 40°C. MS analyses were performed acquiring in MS E mode from 80 to 1500 m/z in positive electrospray ionization mode with a capillary voltage of 3 kV. Data were collected in two channels all of the time: low collision energy (6.0 V), for the molecular ions, and high collision energy (15-40 V), for product ions. The source temperature was set at 110°C. Leucine enkephalin (50 pg/mL) was used as the lock mass (m/z 556.2771) with parameters set to 1 s scan at 20 s intervals. Infusion flow rate for lock mass was 1 µL/min. For unlabeled samples, peak identification, alignment, normalization, and statistical treatment of the data was performed using Progenesis QI software (Nonlinear Dynamics, Waters Corporation, Milford, MA, USA). For 13 Cbiolabeled samples, MassLynx software (Waters Corporation, Milford, MA, USA) was used for data processing and isotopologue labeling distribution analysis.

Heterologous expression of PfPPRD and analysis of CPY glycosylation in Saccharomyces cerevisiae transformants
HA-tagged DFG10 and PfPPRD (optimized for codon usage in yeast) were expressed under the control of the DFG10 endogenous promoter from a centromeric plasmid, pRS416. Empty vector pRS416 was included as a negative control. Saccharomyces cerevisiae strain BY4741 (MATa his3D1 leu2Δ met15Δ ura3Δ) and BY4741-based dfg10Δ strains were grown at 30°C in synthetic defined (SD) medium containing 2% galactose as the carbon source. After two passages (36 to 48 h) in SD medium, cells were harvested when the optical density at 600 nm (OD 600) reached between 0.4 and 1.0. Two OD600 units of cells were harvest for protein analysis and 25 OD600 units of cells were harvested for analysis of polyisoprenoids by LC-HRMS.
Total proteins were extracted as described previously (47). Equal amount of proteins was analyzed using a 12% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to a polyvinylidene difluoride (PVDF) membrane. Mouse anti-CPY (1:5000; Life Technologies) was used as the primary antibody; horseradish peroxidase (HRP)-conjugated antimouse antibody (1:5,000 dilution; Thermo Scientific) together with the Supersignal West Femto substrate (Thermo Scientific) were used to detect the target protein.
For transfection, 20 µg of PfPPRD-TetR-DOZI+HA or PfPPRD-TetR-DOZI plasmids were linearized with EcoRV (NEB), mixed with 60 µg of pUF1-Cas9-guide and precipitated. DNA pellet was dissolved in Cytomix Buffer (37) and combined with fresh red blood cells (RBCs). The DNA/RBC mixture was electroporated using BioRad Gene Pulser Xcell TM system at 0.32 kV, 925 µF. A culture of P. falciparum in schizont stage at high parasitemia was added to the RBCs and parasites were grown in the presence of 0.5 µM aTc (Cayman Chemical). Drug pressure was applied 48 hours post-transfection using 2.5 µg/mL Blasticidin S (BSD) (Sigma). Media was changed daily for the first 7 days and every other day thereafter. P. falciparum cultures were split 1:1 every 7 days to provide fresh uninfected RBCs and to perform DNA extraction for integration analysis.

Growth and morphological assessment of the PfPPRD-TetR-DOZI
In order to assess potential morphological defects due to the reduced expression of PfPPRD, P. falciparum cultures treated with BSD only or BSD+aTc as indicated above were monitored by Giemsa-stained smears every 24 h during 8 days. Potential growth defects were assessed by seeding cultures in a 96 well plate (0.5% early trophozoite stages at 4% hematocrit) and treated with BSD only or BSD+aTc with each condition tested in triplicate. Throughout the course of the experiment, parasites were sub-cultured to maintain the parasitemia between 1-5%. Parasitemia was monitored every 24 hours by flow cytometry using a CytoFLEX S (Beckman Coulter, Hialeah, Florida). A sample of 0.5 µL from each well was stained with Hoechst (Thermo Scientific) at 0.8 µg/mL in PBS and data were analyzed using Prism (GraphPad Software, Inc.). Relative parasitemia at each time point was back calculated based on actual parasitemia multiplied by the relevant dilution factors.

Statistical analysis
The t-test and Benjamini and Hochberg procedure were used to analyze differences between different conditions, and a false discovery rate of 0.01 was used to identify statistically significant differences.     showing that for WT only DOH (15)(16) are detected while in dfg10D mutant strain DOH is accompanied by their respective POH (15)(16). Dfg10D mutant strain transformed with PfPPRD showed considerable reduction of POH 15 and 16.

Figure 6. Successful generation of a P. falciparum PfPPRD-TetR-DOZI conditional knockdown. A)
Schematic of plasmids used and integration of the linearized TetR-DOZI plasmid with 10x aptamer system into PfPPRD gene. The endonuclease cas9 promotes a double-stranded break in the genomic locus while the repair plasmid provides a homology region at the C-terminus of PfPPRD (PfPPRD-C-t) for double crossover homologous recombination to introduce a 3′-untranslated region (UTR) RNA aptamer sequence that binds a tetracycline repressor (TetR) and development of zygote inhibited (DOZI) fusion protein to generate TetR-DOZI conditional system. B) PCR analysis confirming integration at the recombinant locus. Wild type parasites were detected using primers P8 and P2 (Table S1)


extfieldsofstudy: Physics Medicine

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The modular structure of α/β‐hydrolases

The α/β‐hydrolase fold family is highly diverse in sequence, structure and biochemical function. To investigate the sequence–structure–function relationships, the Lipase Engineering Database (https://led.biocatnet.de) was updated. Overall, 280 638 protein sequences and 1557 protein structures were analysed. All α/β‐hydrolases consist of the catalytically active core domain, but they might also contain additional structural modules, resulting in 12 different architectures: core domain only, additional lids at three different positions, three different caps, additional N‐ or C‐terminal domains and combinations of N‐ and C‐terminal domains with caps and lids respectively. In addition, the α/β‐hydrolases were distinguished by their oxyanion hole signature (GX‐, GGGX‐ and Y‐types). The N‐terminal domains show two different folds, the Rossmann fold or the β‐propeller fold. The C‐terminal domains show a β‐sandwich fold. The N‐terminal β‐propeller domain and the C‐terminal β‐sandwich domain are structurally similar to carbohydrate‐binding proteins such as lectins. The classification was applied to the newly discovered polyethylene terephthalate (PET)‐degrading PETases and MHETases, which are core domain α/β‐hydrolases of the GX‐ and the GGGX‐type respectively. To investigate evolutionary relationships, sequence networks were analysed. The degree distribution followed a power law with a scaling exponent γ = 1.4, indicating a highly inhomogeneous network which consists of a few hubs and a large number of less connected sequences. The hub sequences have many functional neighbours and therefore are expected to be robust toward possible deleterious effects of mutations. The cluster size distribution followed a power law with an extrapolated scaling exponent τ = 2.6, which strongly supports the connectedness of the sequence space of α/β‐hydrolases.

Introduction a/b-hydrolases represent a rapidly growing enzyme family with a common fold and a similar active site. The a/b-hydrolase fold consists of a central b-sheet packed between two layers of a-helices [1]. The catalytic triad consists of a nucleophile (serine, aspartate or cysteine), a histidine, a catalytic acid (aspartate or glutamate), and two or three amino acids which form the oxyanion hole. In addition to this conserved, catalytically active core domain, most a/b-hydrolases contain further modules: a lid, a cap, an N-terminal domain, or a C-terminal domain. Despite the high structural similarity of the core domain and the catalytic machinery, a/b-hydrolases show a high sequence diversity and include enzymes with a broad variety of catalytic activities such as acetylcholinesterases, acyltransferases, amidases, dehalogenases, dienelactone hydrolases, epoxide hydrolases (EH), esterases, hydroxynitrile lyases, lipases, peroxidases, proteases and thioesterases [2][3][4][5]. It would have been expected that enzymes with different catalytic activities are separated in sequence space and form separate subfamilies. However, the catalytic activities are overlapping rather than clearly separated in sequence space. Many a/bhydrolases have multiple catalytic activities, and single amino acid positions discriminate between activities such as lipase and amidase [5]. The exchange of single amino acids switched a lipase into an EH [6] or a catalyst for Michael additions [7], a hydrolase into an acyltransferase [8], an esterase into an amidase [9,10], an esterase into an EH [11] and an esterase into a hydroxynitrile lyase [12].
Lipases and other a/b-hydrolases are widely used in biocatalysis because of their high catalytic activity, their regio-and stereoselectivity, their stability in nonpolar media and their ability to catalyse alcoholysis, esterification and transesterification reactions [13,14]. Metagenomic screenings have discovered an ever increasing stream of new sequences [15,16]. Therefore, there is a growing need for the classification of newly discovered sequences into protein families and for the prediction of biochemical and biophysical properties from sequence information [17]. Protein family databases on a/b-hydrolases are versatile tools to systematically compare protein sequences, to annotate functionally relevant amino acids and to assign new sequences to already known subfamilies. Protein family databases such as the Lipase Engineering Database (LED) [18], the database of the a/b-hydrolase fold superfamily ESTHER [19] and the a/b-Hydrolase Fold 3DM Database ABHDB [20] provide a large collection of sequences and structures of a/b-hydrolases, group them into protein families, perform multiple sequence alignments, and derive sequence profiles.
Although sequence alignment allows for a systematic analysis of close homologues, it misses structural and functional relationships between remote homologues. Therefore, a classification based on structural and functional properties is needed. The systematic comparison of the amino acids forming the oxyanion hole allowed to assign a/b-hydrolases to two classes, the GX-and GGGX-types [18]. In GX-types, the first part of the oxyanion hole is formed by the backbone N-H of an amino acid (X) in a GX motif, whereas in GGGX-types it is formed by the backbone N-H of the third glycine in a GGGX-motif. Later, a third type of oxyanion hole (Y-type) was discovered [21]. In Y-type hydrolases, the oxyanion hole is formed by the sidechain of a bulky amino acid, mainly tyrosine or aspartate. The classification by GX-and GGGX-types has high predictive value: in contrast to GX-type, most of the GGGX-types are active toward tertiary alcohols [22]. A systematic comparison of the structures of EHs revealed the modular structure of this subfamily of a/ b-hydrolases with a high similarity within the core domains and within the caps [23]. Cytosolic and microsomal EHs mainly differ by the length of a single loop inside the cap and by their N-terminal domains. Because of their modular structure, the global sequence similarity is misleading, resulting in an overestimation of diversity. Using this classification, the structures of core domains and caps of all EHs and the cap loops of most of the EHs could be reliably modelled [23]. Thus, modularity is key to sequence classification, structure modelling and prediction of function. This analysis was extended to the families of haloalkane dehalogenases and prolyl iminopeptidases, which are homologous to EH despite their different catalytic activities [24]. In the meantime, additional domains were identified in other hydrolases, and their functional role was discussed, such as the N-terminal b-propeller domain in acyl aminoacyl peptidases [25], the role of subdomains in the catalytic behaviour of lipases and acyltransferases [26], the stabilizing N-terminal domain in the murine liver EH (MLEH) [27] or the importance of the C-terminal domain in colipase binding of pancreatic lipases [28].
In this paper, we analysed the architectures of all known a/b-hydrolase structures. The modular structure of a/b-hydrolases can be described as a combination of three core domains (GX, GGGX and Y-types) with caps, N-terminal or C-terminal domains. In addition, a mobile element (the lid) is located at five different positions in lipases. For the analysis, the LED was updated and now contains 280 638 sequences and 1557 structures (https://led.biocatnet.de).

Update of the LED
The LED (https://led.biocatnet.de) was updated starting from the previous version (release 3.0, December 2009) which contained 24 783 sequences from 18 585 proteins and 1117 structures. All structures from this release together with newly identified structures of MHETases [29] were clustered. The resulting centroids were then used as seed sequences (query sequences) for a BLAST search in the NCBI non-redundant protein database and in the Protein Data Bank (PDB), which led to the identification of more than 450 000 putative a/b-hydrolase sequences. One homologous family was created for every centroid sequence, and sequence homologues were added. The centroids of each homologous family and the unclassified structures were analysed visually, revealing a similar core structure which contains the active site, the oxyanion hole residues, and 10 additional modules that can be attached to the core domain: a lid between b-strands b +1 and b +2 , b À1 and b 0 , b À4 and b À3 , b +3 and b +4 , or between the N terminus and b -3 , a single cap, a double cap, an N-terminal cap, two N-terminal domains or a C-terminal domain. These additional modules determine the architecture of the a/b-hydrolase and interestingly some of these elements, such as the lid between b +3 and b +4 and the lid between the N terminus and b À3 , only occur in combination with a C-terminal domain (Fig. 1). One superfamily was created for each architecture. The newly classified sequences were clustered with a similarity threshold of 60%, and new homologous families were created and assigned to their respective superfamilies. Superfamilies were named according to the architecture, whereas the homologous families were named according to their centroid sequence, and groups were formed by the oxyanion hole types (GX, GGGX and Y). Similarly, the superfamilies were assigned to five groups (core, lid, cap, one additional domain or two additional domains). The update resulted in a new version of the LED, which comprises 280 638 individual sequence entries assigned to 198 844 protein entries by a threshold of  98% sequence identity and contains 1557 protein structures in 2772 homologous families and 12 superfamilies. The largest superfamily in the current version of the LED is the N-terminal domain superfamily (GX-and Y-types), containing 25% of the a/b-hydrolases, followed by the single cap superfamily (GX-, GGGX-and Y-types, 23%) the N-terminal cap family (GX-and GGGX-types, 21%), and the core domain family (GX-and GGGX-types, 13%). The smallest protein in the LED with known structure is a lipase of Bacillus subtilis (LED sequence ID 60, PDB-ID: 2QXT and 2QXU) with a length of 179 amino acids and a molecular weight of 19 kDa, the largest protein a dipeptidyl peptidase 8 (LED sequence ID 456274, PDB-ID: 6EOP) with a sequence length of 900 amino acids and a molecular weight of 104 kDa.
Of each homologous family, one protein with available structure was visually analysed, and the catalytic triad, the oxyanion hole residues and structural modules such as lids, single caps, double caps, N-terminal caps, N-terminal domains and C-terminal domains were annotated. The annotation information was then transferred from the centroids to the other members of a homologous family based on a multiple sequence alignment. In total, 162 737 sequences were annotated (58% of all sequences of the LED). Based on all annotation entries in the LED, the lengths of the additional modules and domains were analysed. Lids are the shortest modules with a length of 20 AE 7 amino acids (mean AE standard deviation), N-terminal caps and the N-terminal part of the double caps are similar in size (40 AE 8 and 53 AE 14 amino acids respectively), single caps are larger (75 AE 18 amino acids), and the additional C-and N-terminal domains are largest and vary considerably in size (367 AE 129 amino acids).

Cluster size distribution
Pairs of homologous sequences were derived for 280 638 sequence entries from the updated LED (release 4.0). The sequences were assigned to different clusters (communities), by setting a threshold of global sequence identity to 60%, 70%, 80% or 90%. The number of clusters N(s) with a cluster size s decreased with increasing s following a power-law distribution N (s)~s Às (Fig. 2). Logarithmic histograms were formed for s ≥ 2 and s ≤ 10, s ≥ 11 and s ≤ 100, . . ., s ≥ 1001 and s ≤ 10 000 sequences [30]. s h was determined from the slope of the histograms as 0.8, 1.0, 1.2 and 1.4 at a threshold of global sequence identity of 60%, 70%, 80% and 90%, respectively (Table 1), and the underlying s was modelled as described previously [30]. For comparison, s was also determined by linear regression of the actual distribution for s ≤ 100 (Fig. S1), and both methods resulted in the same values of s, ranging from 1.7 to 2.3 between 60% and 90% global sequence identity. s was then extrapolated to 100% global sequence identity, yielding s 100 = 2.6 ( Fig. S2), which is identical to the value derived previously for a/b-hydrolases [30].

Degree distribution
As further property of the network topology, the number of neighbouring sequences (degrees), was analysed. Neighbouring sequences were defined by a threshold of global sequence identity of 95%. The number N(n) of sequences having n neighbours followed a power law distribution N(n)~n Àc . Although many sequences had few neighbours, a small number of sequences had a high degree and were located in a hub region. Linear regression was performed for n ≤ 80 resulting in a scaling exponent c = 1.4 ( Fig. 3), which is similar to the values of c between 1.1 and 1.3 determined previously for five different protein families [31]. All sequences in the hub region, (n ≥ 300) belong to a single homologous family, the protease 2 homologous family number 1011, which are Y-type a/b-hydrolases from the N-terminal domain superfamily number 8 (Table S1).

Modular structure of a/b-hydrolases
One representative a/b-hydrolase structure for each architecture and oxyanion hole type was visually analysed. The structures of centroids were selected if they were fully resolved and had a resolution of at least 3 A. Otherwise, a structure with high similarity to the centroid was selected. All a/b-hydrolases have a similar core structure, the a/b-hydrolase fold, which contains the catalytic triad and the oxyanion hole residues (Fig. 1). The b-strands of the central b-sheet were numbered starting from b 0 , the b-strand preceding the nucleophilic elbow, where the catalytic serine is located. All b-strands in the direction of the N terminus were assigned negative numbers (. . ., b À3 , b À2 , b À1 ), whereas the b-strands in the direction of the C terminus were assigned positive numbers (b +1 , b +2 , b +3 ,. . .) (Figs S3-S9). As an additional module, a/b-hydrolases might contain a mobile lid consisting of one or two a-helices. The lid can only be unambiguously assigned if open and closed conformations of the same or two homologous proteins have been crystallized. Therefore, in the absence of both conformations, a/b-hydrolases with a lid might be misclassified as core domain a/b-hydrolases. The lid is located at different positions within the central b-sheet: between b-strands b +1 and b +2 , b À1 and b 0 , b À4 and b À3, b +3 and b +4 , or between the N terminus of the protein and b À3 . In contrast to the mobile lid, a cap is immobile and consists of more than two ahelices which are stacked on top of the core structure. Three cap arrangements were found: a single cap located between b-strands b +1 and b +2 , a double cap where an additional N-terminal cap is stacked on top of the single cap, and an architecture with an N-terminal cap only. In addition, a/b-hydrolases might contain an N-terminal domain with a b-propeller fold or a Rossmann fold, or a C-terminal domain with a b-sandwich fold. The N-terminal Rossmann fold domain and C-terminal domains can be combined with a cap or a lid, respectively. Thus, in total 12 different architectures were identified (Fig. 1). In addition, a/b-hydrolases can be distinguished by their oxyanion hole ( Table 2). Ten of the 12 architectures contain GX-types, but only four architectures contain GGGX-types (core, core with lid  between b À3 and b À4 , core with a single cap or with an N-terminal cap) and three architectures contain Ytypes (core with a single cap, core with an N-terminal b-propeller domain or core with a C-terminal b-sandwich domain).

Core domain a/b-hydrolases
The core domain of a/b-hydrolases is composed of up to 13 b-strands forming a central parallel b-sheet. The b-strands alternate with varying numbers of a-helices, and the b-sheet is packed between two layers of a-helices. The central b-strand b 0 is connected to the subsequent a-helix by a sharp turn, the nucleophilic elbow, which harbours the catalytic serine and one of the oxyanion hole residues (Fig. S3). a/b-Hydrolases consisting only of the core domain were found in the GXtypes (29 712 sequences, representative structure: lipase A from B. subtilis, BSLA) and in the GGGX-types (5376 sequences, representative structure: carboxylesterase from Lactobacillus plantarum, LPC). As of now, no Y-type a/b-hydrolases were found that consist exclusively of the core domain ( Table 2). The active site of core domain a/b-hydrolases is completely exposed to the solvent (Fig. S10A). In contrast to GX-types, additional a-helices restrict the substrate access to a specific direction in most GGGXtypes, thus contributing to substrate specificity (Figs S11A and S12A), because GGGX-types are larger (200-500 amino acids, 8-13 b-strands) than GX-types (180-300 amino acids, 5-8 b-strands) and thus have more helices close to the substrate access tunnel that can shield the active site (Table S3).
Lids as an opening and closing mechanism for the active site  Table 2). GGGX-type a/b-hydrolases can have a lid between b À4 and b À3 (9079 sequences, representative structure: lipase from Candida rugosa in open and closed conformation, Table 2. Basic architectures of a/b-hydrolases. Representative structures were chosen for the different oxyanion types and architectures. Abbreviations of protein names can be found in Table S2. In addition to the architecture, the position of the additional module in the central b-sheet is indicated. The letter in brackets following the protein name indicates the amino acid of the active site nucleophile. - CRL open and CRL closed ) ( Fig. S4C and Table 2). No Ytype a/b-hydrolases with a lid has been found, yet. Regardless of where the lid emerges, its position in respect to the active site is almost identical. In its closed conformation, it covers the active site from the N-terminal side of the protein and moves towards the N terminus of the protein upon opening (Figs 1B-D, S10 and S11B-D). When the lid is in the closed conformation, the active site residues are not visible and the contact area of the lid surrounds the active site completely. This suggests that the lid covers the active site in the closed conformation, making the active site inaccessible to solvent or substrate molecules. Upon opening of the lid, the contact area of the lid moves away from the active site, thus making the active site fully accessible to the solvent and thereby allowing substrate molecules to bind to the active site (Figs 4 and 5).

Caps covering the active site
Caps are immobile elements, which cover the active site either partly or completely. a/b-Hydrolases can have single caps, N-terminal caps or double caps added to the core domain ( Fig. 1E-G). Single caps consist of three or more a-helices on top of the proteins (Figs S5A and S11E). N-terminal caps consist of two or more long a-helices formed by the N terminus folding back over the active site (Figs S5B and S11F).
In a/b-hydrolases with a double cap, two caps are stacked on top of the active site. The lower cap is similar to the single cap, the upper cap to the N-terminal cap (Figs S5C and S11G). a/b-Hydrolases with a single cap were found as GXtypes (61 596 sequences, representative structure: haloalkane dehalogenase from Xanthobacter autotrophicus, XAHD), GGGX-types (2581 sequences, representative structure: HsaD from Mycobacterium tuberculosis, MTH), and Y-types (22 sequences, lipase A from Candida antarctica, CALA) ( Table 2), and is the only architecture that can be found for all oxyanion hole types. The position of the single cap is conserved and is always located between strand b +1 and b +2 (Fig. S5A). The active site of Y-type a/b-hydrolases with a single cap is completely covered by the cap, thereby restricting access to the active site. In GX-types, the active site is mostly covered, however, the catalytic triad histidine is partially exposed to the solvent. In GGGX-types, the entrance to the active site from the top is completely covered by the cap, but there is a narrow tunnel between the cap and the core domain which leads to the active site (Figs 6A and S10E).
In N-terminal caps, the helices restrict the access of solvent or substrate molecules to the active site (Figs 6B, S10F and S122B). a/b-Hydrolases with an N-terminal cap could be found as GX-types (585 sequences, representative structure: cutinase from Trichoderma reesei, TRC) and as GGGX-types (57 900 sequences, representative structure: esterase from Pyrobaculum calidifontis, PCE), but not as Y-types. In GX-types, the N-terminal cap blocks the active site completely, suggesting that the substrate access to the active site is restricted (Fig. S13A). In contrast, in GGGX-types, the cap does not block the active site completely, resulting in a narrow tunnel in the side of the protein leading towards the back of the central helix of the active site, which might allow substrate access (Fig. 6B).
Until now, double caps have only been found in GX-type a/b-hydrolases (14 874 sequences, representative structure: EH from Streptomyces carzinostaticus, SCEH) ( Table 2). The combination of two caps covers the active site completely without leaving a tunnel between the caps (Figs S10G, S12C and S13B).
a/b-hydrolase fusion proteins with additional N-or Cterminal domain The modules described so far are relatively small and interfere with the entrance to the active site. Another group of a/b-hydrolases displays additional domains that are attached to their N-or C terminus (Fig. 1H, I). a/b-Hydrolases with an additional N-terminal bpropeller domain were found for GX-types (15 436 sequences, representative structure: acylaminoacyl peptidase from Aeropyrum pernix K1, APAP) and Y-types (53 669 sequences, representative structure: human dipeptidyl peptidase IV [DPPIV]) ( Table 2). The bpropeller domain consists of seven to eight blades, each blade is formed by a four-stranded b-sheet. (Figs 7A and S6). The b-propeller domain is stacked on top of the core domain, and the blades are arranged to form a channel through the middle of the propeller towards the active site (Fig. S10I). In Ytypes, the propeller domain does not block the active site completely, suggesting there might be an additional substrate access between the propeller and the core domains (Figs 7A and S13C).
a/b-Hydrolases with an additional C-terminal b-sandwich domain were found in the Y-types only (13 624 sequences, representative structure: cocaine esterase from Rhodococcus sp. MB1, RCE) and not in GX-types or GGGX-types (Figs 7B, S7 and Table 2). In the CATH database, the C-terminal b-sandwich domain is annotated as galactose-binding domain-like, CATH superfamily 2.60.120.260 [32]. The C-terminal b-sandwich domain does not interfere with the active site and is located at the side of the protein. Therefore, a/b-hydrolases with a C-terminal b-sandwich domain have a freely accessible active site (Figs 7B, S10H and S13D).
Proteins with two additional modules a/b-Hydrolases with more than one additional module are rare and only found for GX-types (Fig. 1J,L): a/bhydrolases with a C-terminal domain and a lid  Although the opening/closing transition and the size of the lids are similar, the location of the lid differs. In HPL/HPLRP1, it is located between strands b +3 and b +4 , whereas in PSML it is located between the N terminus and b À3 (Fig. S8A,B). Interestingly, although the lids emerge from different locations within the central b-sheet, they are situated in a similar position covering the active site from the C terminus of the protein, rather than from the N terminus as observed for other a/b-hydrolases (Fig. 1). The N-terminal domain of MLEH consists of a Rossmann fold formed by alternating a-helices and bstrands, annotated by CATH classification as HAD superfamily/HAD-like domain, or CATH Superfamily 3.40.50.1000 [32]. Other than the N-terminal b-propeller domains and similar to the C-terminal b-sandwich domains, this N-terminal Rossmann fold domain is attached to the side of the protein and does not interfere with substrate access to the active site (Figs 8B and S12D). The cap of MLEH is comparable to other single caps and is located between b +1 and b +2 (Figs 8B and S9). Although it covers the active site, there seems to be a tunnel underneath the cap, which allows access of substrates to the active site (Figs 8B and S10L).

Similarity of the N-and C-terminal domains to other proteins
In order to analyse whether the N-and C-terminal domains in superfamilies 8-12 are similar to domains in other proteins, one representative protein from each of these superfamilies and each oxyanion hole type was selected: from the N-terminal domain superfamily number 8, the GX-type A. pernix acylaminoacyl peptidase (APAP) and the Y-type human DPPIV, from the C-terminal domain superfamily number 9, the Y-type Rhodococcus sp. cocaine esterase (RCE), from the lid (b +3 and b +4 ) and C-terminal domain superfamily number 10, the GX-type HPL, from the lid (N-terminal) and C-terminal domain superfamily number 11, the GX-type Pseudomonas sp. MIS38 lipase (PSML), and from the cap and N-terminal domain superfamily number 12, the GX-type human epoxide hydrolase (HEP) ( Table S4).
The N-terminal b-propeller domains of DPPIV and APAP are structurally similar, as well as the b-sandwich domains of RCE and HPL. All a/b-hydrolase core domains are similar to each other. In addition, they are similar to the additional N-terminal Rossmann fold domain of HEP (Fig. S14).
In a second step, the PDB was searched for domains with structural similarity to the N-or C-terminal domains (https://doi.org/10.18419/darus-458).
The search with the b-propeller domains of DPPIV and APAP resulted in 103 and 173 structurally similar proteins, respectively, which consist of a single propeller, two fused propellers, or fusion proteins of a bpropeller and other domains. They cover a broad range of biological functions, such as transcription and translation initiation factors, DNA damage-binding proteins, elongation complexes, export factors, splicing factors, ribosomal proteins, proteins involved in cell cycle, apoptosis, and intracellular transport, and also b-propeller lectins, although the carbohydrate-binding site does not seem to be conserved. The search with the Rossmann fold domain of HEP resulted in 154 hits of structurally similar proteins. Because of its similarity to the a/b-hydrolase core domain, 54% of all hits belonged to a/b-hydrolases, mostly single cap a/b-hydrolases from LED superfamily 5. All other hits were mainly proteins consisting of a single Rossmann fold domain, two fused Rossmann fold domains or fusion proteins of Rossmann fold with other domains. They cover a broad range of enzymatic functions, such as phosphatases, haloacid dehalogenases, phosphoglucomutases, reductases and glycosyltransferases, but also regulators of transcription or replication.
The search with the b-sandwich domains of RCE, PSML and HPL resulted in 41, 35 and 10 proteins, respectively, which consist of a single b-sandwich or are fusion proteins of b-sandwich domains with mainly ahelical domains. Proteins with structural similarity to HPL cover a broad range of annotations such as lipoxygenases, glucanases, xylanases, the GTPase Rab6 and carbohydrate-binding modules. Proteins with structural similarity to RCE are involved in DNA repair, flagellar biosynthesis or receptor binding, but also lectins and enzymes involved in carbohydrate degradation, such as xylanases, glycoside hydrolases, mannanases and endoglucanases. Although the structural similarity suggests a carbohydrate-binding function, the carbohydrate-binding site of lectins is not present in the b-sandwich domains of RCE and HPL. Proteins with structural similarity to PSML include hemagglutinin, adhesins, toxin A and antifreeze proteins.

Naming conventions
Despite their high variability in sequence and structure, a/b-hydrolases can be assigned to a small number of classes. Two criteria were used: the sequence motif of the oxyanion hole and the presence of lids, caps and N-or C-terminal domains. However, there is no consistent naming of these structural modules in literature, especially for the definition of lids and caps. The definition of a 'cap' as a large immobile module covering the active site is widely accepted [33][34][35][36]. However, in some publications it is referred to as 'lid' [2,37,38]. Similarly, small mobile elements covering the active site are mostly referred to as 'lid' [35,[39][40][41][42][43], but they might also be called 'flap' [44][45][46][47]. To improve communication about a/b-hydrolases, we suggest the following naming conventions: A 'lid' is a small mobile structure comprised of one or two a-helices, which can undergo a conformational transition, thereby opening and closing the entrance to the active site. Lids are located between b-strands b À3 and b À4 , b 0 and b À1 , b +1 and b +2 , b +3 and b +4 or between the N-terminus and b À3 .
A 'cap' is an immobile module and consists of three or more a-helices, which cover the active site. Caps are located between strands b +1 and b +2 . In proteins with double caps, a second cap consisting of two or three a-helices is located at the N terminus. This second cap is similar to N-terminal caps. These are formed by two or more a-helices emerging from the N terminus of the protein, which fold over the active site and thereby cover it.
N-terminal domains have either a b-propeller fold or a Rossmann fold. The C-terminal domains have a b-sandwich fold. These additional domains can be located on top of the core domain covering the active site, on its side or separated by a long loop.
Although caps and the N-terminal or C-terminal domains can be easily identified in protein structures, the unambiguous identification of one or two a-helices as lid requires the presence of an open and a closed conformation, such as in C. rugosa lipase (PDB-ID: 1CRL and 1TRH), Rhizomucor miehei triacylglyceride lipase (PDB-ID: 3TGL and 4TGL), or Pseudomonas sp. MIS38 lipase (PDB-ID: 2Z8X and 2ZVD).
The suggested classification by oxyanion hole motif and by the presence of lids, caps and N-or C-terminal domains is based on the analysis of protein structures. However, a few putative a/b-hydrolases with known structure could not be assigned because of deviations of their oxyanion hole sequence from the conserved motifs, which did not allow us to assign them to GX-, GGGX-or Y-types: The N-acyl homoserine lactone degrading enzyme (PDB-ID: 5EGN) and a hydrolase from Pseudomonas aeruginosa PA01 (PDB-ID: 3OM8) had a cap, but their oxyanion hole motifs were PF and SI respectively. Similarly, the putative dienelactone hydrolase from Klebsiella pneumoniae (PDB-ID: 3F67) and an uncharacterized protein from Escherichia coli (PDB-ID: 4ZV9) are core domain a/bhydrolases, however, their oxyanion hole motif was EX.
In addition, the database update resulted in % 170 000 discarded sequences which could not be assigned to one of the superfamilies because of their low sequence similarity to proteins with known structure, which made it impossible to decide about the presence of modules.

Substrate access
The knowledge of the structural elements of a protein allows a prediction about the access of substrate to the active site. For a/b-hydrolases that consist only of the core domain, the active site is fully exposed to the solvent. This was already shown for the lipase from B. subtilis, the cutinase from Fusarium solani or the carboxylesterase from Pseudomonas fluorescens [48][49][50]. In a/b-hydrolases with a lid, the substrate enters between core and lid. In the closed conformation, the active site is covered by the lid, but becomes accessible to the substrate upon a conformational transition of the lid to an open conformation, as shown for the lipases from C. rugosa, P. aeruginosa or Rhizopus niveus [51][52][53]. In contrast, the cap covers the active site. In order to access to the active site, a substrate molecule has to pass through a tunnel located between the core and the cap, such as in the HEP, the prolyl aminopeptidase from Serratia marcescens, or the C-C hydrolase from M. tuberculosis [54][55][56], or through a tunnel in the cap, such as in the fluoroacetate dehalogenase from Rhodopseudomonas palustris or the hydroxynitrile lyase from Hevea brasiliensis [57,58]. Access to the tunnel is controlled by gatekeepers such as the sidechain of Leu262 in the haloalkane dehalogenase from X. autotrophicus [59]. Engineering of the tunnel can result in changes of substrate specificity [60]. Interestingly, in GGGX-type proteins with a single cap or an N-terminal cap, the active site is partially solvent accessible, which, however, might be a crystallization artefact. For Y-type a/b-hydrolases with an N-terminal b-propeller domain, substrate access is under discussion. For dipeptidyl peptidases, the substrate was shown to enter the active site through a side opening, along the interface between N-terminal and core domain [61,62]. The function of the tunnel in these proteins is not clear, but was suggested to aid the release of product from the active site [63,64]. For prolyl oligopeptidases, which are also Y-type a/b-hydrolases with an N-terminal b-propeller domain, a crystal structure with a tilted propeller domain was identified, resulting in a large opening between the core domain and the propeller domain, similar to dipeptidyl peptidases [37]. It has therefore been discussed whether substrate molecules enter prolyl oligopeptidases through the tunnel of the propeller domain [65,66] or through the entry site that opens upon conformational change of the proteins [37,67,68].

Sequence space
Despite the large number of 280 638 sequence entries in the updated LED, we still know only a tiny fraction of the whole extant sequence space of a/b-hydrolases [69]. The limited coverage of sequence space and the even lower coverage of structure space might explain why GX-types were found in combination with most modules and multiple lid positions, but only few combinations and lid positions were found for GGGX-and Ytypes. However, the analysis of the sequence network confirmed previous results on the properties of protein sequence space. The scale-free distribution of the number of neighbours of a protein sequence demonstrates the existence of a few hubs and a large number of loosely connected sequences, as found previously for five different protein families. The scaling exponent of the a/b-hydrolase sequence network (c = 1.4) is similar to other protein families which had scaling exponents between 1.1 and 1.3 [31]. The members of the homologous family 1011 (annotated as 'protease 2' or 'oligopeptidase B') had the highest number of neighbours (Table S1) and thus formed the largest hub region of the a/b-hydrolase network. Since hub sequences have many functional neighbours, they have proven to be highly evolvable with respect to robustness towards mutations [70]. Because mutations might readily induce new functions, sequences with a large number of neighbours are promising starting points in directed evolution experiments.
A second property of the a/b-hydrolase network, the scale-free cluster size distribution, is also similar to other protein families [30]. The extrapolated scaling exponent (s 100 = 2.6) was in the range between 2.3 and 3.3 which was previously determined for six protein families [30] and indicates percolation, that is, connectedness, of protein sequence space, despite the fact that extant sequence space covers only a tiny fraction of theoretic sequence space, yet. The connectedness of protein sequence space implies that various evolutionary pathways between two homologous sequences are possible, favouring substrate ambiguity or promiscuity [71][72][73].

Structure space
Although global sequence similarity is a widely used measure of the relationship between proteins, it is misleading. In addition to exchanges, insertions and deletions of single amino acids, there is a second evolutionary mechanism: the recombination of structural modules or domains, which has been observed in many protein families, such as thiamine diphosphatedependent enzymes [74] or glycoside hydrolases and carbohydrate-binding modules [75]. Although the identification of structural and functional domains and modules in a protein structure might be challenging [76], many proteins which are distant considering their global sequence and structure share highly similar fragments, which resulted in the view that protein structure space is continuous rather than discrete [77].
The sequence similarity between a/b-hydrolases is generally low and their similarity relies on their structure [3]. The similarity of domains can be hidden by the global similarity, especially if different modules are recombined and exist in different orders [74]. Although the global structural similarity is quite low or undetectable between some a/b-hydrolases with additional C-and N-terminal domains, the structural similarity of the core domains to each other is remarkable. This is different for the additional modules and most of them are structurally quite diverse. However, there are exceptions, such as the b-propeller domain of Y-type and GX-type a/b-hydrolases. These are structurally very similar and so are the core domains, pointing to coevolution. Interestingly, also the core domain and the additional C-terminal b-sandwich domain of HPL and RCE show significant structural similarity. Interestingly, RCE is a Y-type with an additional C-terminal b-sandwich domain, whereas HPL belongs to the GXtypes and has an additional C-terminal b-sandwich domain and a lid attached. This suggests that existing modules can be exchanged or newly added to proteins. In fact, the addition or deletion of N-and C-terminal domains to proteins was described as one of the most frequent domain rearrangements observed [78]. The substitution of domains inside a protein was, however, shown to be rare [79], which raises the questions, whether lids and caps can also be exchanged between proteins. Interestingly, the fold of the additional Nand C-terminal domains also occurs in several other proteins with very different functions. Although the sequence similarity between these modules is very low, they share structural similarity, showing how evolution reuses suitable folds for completely new functions.
Continuous protein space is supported by another very interesting finding in the LED that proteins could clearly be assigned to one of the homologous families although they are enzymatically inactive and display no hydrolase activity. One such an example is the Gibberellin receptor gibberellin insensitive dwarf1 (GID1) (PDB-ID: 2ZSI and 3ED1). This protein can clearly be assigned to GGGX-types with an N-terminal cap, but they miss the catalytic histidine. This is interesting, because the histidine is the most conserved residue of the catalytic triad of a/b-hydrolases [3]. Indeed, it was shown that although GID1 is similar to hormone-sensitive lipases, it does not display any hydrolytic activity [80][81][82]. Another interesting example are neuroligins, for example, the extracellular domain of neuroligin 2A from mouse (PDB-ID: 3BL8) or neuroligin-1 from rat (PDB-ID: 3BIW). Both can clearly be assigned to the GGGX-type a/b-hydrolases with an additional lid, however, they are missing the catalytic serine and instead show a GXGXG motif, which leads to an enzymatically inactive protein [83,84]. This demonstrates how the structure of a/b-hydrolases can be exploited and used for other functions.
A few years ago, plastic-degrading enzymes were discovered as an interesting possibility for the bioconversion of polyethylene terephthalate (PET) [85]. These so called PETases are able to convert PET to mono-(2-hydroxyethyl) terephthalate (MHET), which is then hydrolysed by MHETases to terephthalate and ethylene glycol [29]. PETases and MHETases are members of the a/b-hydrolase family [29,85] and can be found in the LED. PETases are GX-types, whereas MHETases are GGGX-types, but both are core domain a/b-hydrolases. MHETases were suggested to have a lid that confers substrate specificity [29], however, the domain specified by Palm and colleagues only shields, but does not cover the active site. Furthermore, it consists of 8 a-helices and 242 amino acids and thus is too big to be classified as a lid according to the naming convention suggested in this paper. In the LED, PETases and MHETases are found in superfamily number 1. There are 1099 PETases and homologues thereof combined in homologous family number 49 and 13 MHETases and homologues thereof in homologous family number 2923. Therefore, the LED can aid in identifying possible PETases and MHETases. It further allows to find related proteins that could be used in protein engineering to develop highly active plastic-degrading enzymes.

Setup of the Lipase Engineering Database (LED)
The sequences of 1117 a/b-hydrolases with structure information in the previous version of the LED (release 3.0, December 2009) were clustered using USEARCH (version 11.0.667) with an identity threshold of 90%, resulting in a list of representative sequences (named centroids) [86]. The structures of the centroids were checked visually for their architecture using the PyMOL Molecular Graphics System (version 1.8.0.5, Schr€ odinger; LLC, New York, NY, USA). Structures lacking the a/b-hydrolase fold were discarded, and the remaining centroid sequences were reclustered by USEARCH with an identity threshold of 60% to reduce the number of queries for later BLAST searches. In addition, the sequences of three recently identified structures of MHETases, which also belong to the family of a/b-hydrolases, were added to the centroids [29]. Each centroid served as a seed sequence of a homologous family and was assigned to a superfamily [87], which was named by its respective architecture. By using the centroid sequences as query, homologous proteins were searched in the NCBI non-redundant protein database and the PDB by BLAST searches with an E-value cut-off of 10 À10 [88,89], resulting in more than 450 000 putative a/b-hydrolase sequences. A threshold of 98% global sequence identity was used to assign individual sequences to proteins. Sequences were sorted into homologous families at a global sequence similarity threshold of 60%. For each of the homologous families, a profile hidden Markov model (HMM) was created using HMMER 3.1b2 (http://hmmer.org/). Iterative HMM searches with these profiles in the remaining unassigned sequences were performed with decreasing E-values from 1.5•10 À5 to 5•10 -10 , and the hits were assigned to the respective superfamily after a visual inspection of their architecture. The unassigned sequences were excluded from further analyses. The newly assigned sequences were clustered with an identity threshold of 60% using USEARCH. For each cluster containing more than nine sequences, a new homologous family was created in the respective superfamily. All sequences in a superfamily, which were not assigned to a homologous family, were summarized in the 'singleton family'.

Visual analysis of a/b-hydrolase structures
The PyMOL Molecular Graphics System, (version 1.8.0.5, Schr€ odinger; LLC) was used for the visual analysis of a/bhydrolases and the generation of figures. Contact areas of the modules with the core domains were calculated using a PyMOL script by Martin Christen, in which the contact radius was increased to 6 A (Martin Christen, 2013, Contact Surface, v.3.0, https://pymolwiki.org/index.php/Contac t_Surface). Explosion graphs were created by disassembling the structures and pulling the modules (lids, caps, N-or Cterminal domains) away the from the core protein.

Analysis of protein sequence networks: clusters and degrees
Pairwise global sequence alignments for the 280 638 sequence entries from the updated LED were used to derive edge weights of pairwise sequence identity for protein sequence networks. Instead of aligning all sequence pairs, the calc_distmx command from USEARCH (version 11.0.667) was used to heuristically determine pairs of homologues, thereby reducing computational effort [86]. Subsequently, the sequence pairs were aligned using the Needleman-Wunsch algorithm implemented in the EMBOSS software suite (version 6.6.0, EMBL-EBI, Hinxton, UK) with gap opening and gap extension penalties of 10 and 0.5 respectively [90,91]. Protein sequence networks were constructed as undirected graphs with edge weights of global sequence identity. Thresholds of global sequence identity were applied to form clusters (communities) of homologous sequences. The number of nodes N(s) of a cluster size s was fitted by a power law N(s)~s Às , and the Fisher exponent s was determined from the slope of the distribution in a log-log plot [30]. Logarithmic histograms were formed for s ≥ 2 and s ≤ 10, s ≥ 11 and s ≤ 100, . . ., s ≥ 1001 and s ≤ 10 000 sequences. The slopes of these histograms, s h , were determined at varying thresholds of global sequence identity. The slopes of the actual distribution, s, were derived by fits of s h against a power-law model distribution as described previously [30]. The number of neighbouring nodes n at a given threshold is called the degree of a node. The number of nodes N(n) having a degree of n was fitted by a power law N(n)~n Àc , and the scaling exponent c was derived from a log-log plot [31]. Distributions of cluster sizes and degrees were analysed by linear fitting via the fitlm function from the Statistics and Machine Learning Toolbox (version 11.5) in MATLAB (version R2019a The MathWorks, Natick, MA, USA).

Annotation of protein sequences
If available, a representative protein with structure information was selected from each homologous family, and the catalytic nucleophile (serine, aspartate or cysteine), histidine and the catalytic acid (aspartate or glutamate) were annotated, as well as the residues forming the oxyanion hole, the lid, single cap, secondary cap (N-terminal part of double caps), N-terminal cap, N-terminal domain and C-terminal domain. For each homologous family, a multiple sequence alignment was generated using Clustal Omega (version 1.2.1) [92]. The annotations were then transferred to the respective positions in the aligned sequences. Thus, the sequences of all homologous families that contain at least one protein structure were annotated in the updated release of the LED (release 4.0).

Structural similarities of the N-and C-terminal domains
To analyse the structural similarities of the N-and C-terminal domains, representative protein structures of each oxyanion hole type from the proteins of superfamilies 8-12 were selected. The structures of the N-and C-terminal domains and the core domains were extracted and saved in separate.pdb files, which were then used to compare the domains by the 'all against all' structure comparison tool on the Dali server [93]. In a second step, the PDB search tool on the Dali server was used to compare the N-and Cterminal domains against all structures in PDB25, a subset resulting from clustering the whole PDB database with an identity threshold of 25%. The Dali server has different measures to determine similarity between proteins. The Z-Score is a measure for structural similarity, where a Z-score > 2 implies significant structural similarities and thus a similar fold [94]. Besides the Z-Score, the output also contains information about the rmsd and the sequence identity.

Supporting information
Additional supporting information may be found online in the Supporting Information section at the end of the article. Table S1. Exemplary hub sequences with their annotated source organisms and descriptions. Table S2. List of the proteins names and abbreviations of the representative structures. Table S3. Sequence length of the representative protein structures retrieved from the PDB. Table S4. Representative structures used for the structural comparison of additional modules to other proteins using the Dali web server. Fig S1. Cluster distributions N(s) with linear regressions for cluster sizes s ≤ 100 at thresholds of global sequence identity of 60% (A), 70% (B), 80% (C) and 90% (D). Fig. S2. Fitted exponents s (dots), derived from fits of the slopes of the histograms, s h , against the model distribution from , determined for different thresholds of global sequence identity. Fig. S3. Topology diagram of the core domain of a/bhydrolases. Fig. S4. Topology diagram of a/b-hydrolases with an additional lid (green). Fig. S5. Topology diagram of a/b-hydrolases with an additional single cap (red), N-terminal cap (pale red) or double cap. Fig. S6. Topology diagram of a/b-hydrolases with an additional N-terminal propeller domain (blue). Fig. S7. Topology diagram of a/b-hydrolases with an additional C-terminal b-domain (blue). Fig. S8. Topology diagram of a/b-hydrolases with two additional domains: an additional lid (green) and Cterminal b-sandwich domain (blue).