Journal of Structural Biology 184 (2013) 182–192

Contents lists available at ScienceDirect

Journal of Structural Biology

journal homepage: www.elsevier .com/ locate/y jsbi

Crystal structure of rat intestinal alkaline phosphatase – Role of crowndomain in mammalian alkaline phosphatases

1047-8477/$ - see front matter � 2013 Elsevier Inc. All rights reserved.http://dx.doi.org/10.1016/j.jsb.2013.09.017

Abbreviations: APs, alkaline phosphatases; EcAP, Escherichia coli alkaline phos-phatase; PLAP, placental alkaline phosphatase; IAP, intestinal alkaline phosphatase;GCAP, germ cell alkaline phosphatase; TNAP, tissue non-specific alkaline phospha-tase; pNPP, p-nitrophenyl-phosphate; rIAP, rat intestinal alkaline phosphatase;cIAP, cynomolgus monkey intestinal alkaline phosphatase; rIAP-Ic, rat intestinalalkaline phosphatase expressed in Sf9 insect cell; rIAP-Ec, rat intestinal alkalinephosphatase expressed in E. coli; TM, triple mutant; Tm, melting temperature; Nag,N-acetyl glucosamine.⇑ Corresponding authors.

E-mail addresses: Kaushik.Ghosh@syngeneintl.com (K. Ghosh), manjunath.ramarao@bms.com (M. Ramarao).

Kaushik Ghosh a,⇑, Debarati Mazumder Tagore a, Rushith Anumula a, Basanth Lakshmaiah a,P.P.B.S. Kumar a, Senthuran Singaram a, Thangavelu Matan a, Sanjith Kallipatti a, Sabariya Selvam a,Prasad Krishnamurthy a, Manjunath Ramarao b,⇑a Applied Biotechnology, Biocon Bristol-Myers Squibb Research and Development Center, Syngene International Ltd., Biocon Park, Bommasandra IV Phase, Jigani Link Road,Bangalore 560099, Indiab Bristol-Myers Squibb India Ltd. and Biocon Bristol-Myers Squibb Research and Development Center, Bangalore 560099, India

a r t i c l e i n f o a b s t r a c t

Article history:Received 2 July 2013Received in revised form 20 September 2013Accepted 21 September 2013Available online 27 September 2013

Keywords:Rat intestinal alkaline phosphatasestructureCrown domainMetal ion requirementsGlycosylation

Intestinal alkaline phosphatases (IAPs) are involved in the cleavage of phosphate prodrugs to liberate thedrug for absorption in the intestine. To facilitate in vitro characterization of phosphate prodrugs, we havecloned, expressed, purified and characterized IAPs from rat and cynomolgus monkey (rIAP and cIAPrespectively) which are important pre-clinical species for drug metabolism studies. The recombinantrat and monkey enzymes expressed in Sf9 insect cells (IAP-Ic) were found to be glycosylated and active.Expression of rat IAP in Escherichia coli (rIAP-Ec) led to �200-fold loss of activity that was partially recov-ered by the addition of external Zn2+ and Mg2+ ions. Crystal structures of rIAP-Ec and rIAP-Ic were deter-mined and they provide rationale for the discrepancy in enzyme activities. Rat IAP-Ic retains its activity inpresence of both Zn2+ and Mg2+ whereas activity of most other alkaline phosphatases (APs) including thecIAP was strongly inhibited by excess Zn2+. Based on our crystal structure, we hypothesized the residueQ317 in rIAP, present within 7 Å of the Mg2+ at M3, to be important for this difference in activity. TheQ317H rIAP and H317Q cIAP mutants showed reversal in effect of Zn2+, corroborating the hypothesis. Fur-ther analysis of the two structures indicated a close linkage between glycosylation and crown domainstability. A triple mutant of rIAP, where all the three putative N-linked glycosylation sites were mutatedshowed thermal instability and reduced activity.

� 2013 Elsevier Inc. All rights reserved.

1. Introduction are largely conserved. Escherichia coli AP (EcAP) is one of the most

Alkaline phosphatases (APs), a class of hydrolase (EC 3.1.3.1)common to all organisms, are involved in the hydrolysis of phos-phate monoesters under alkaline conditions. They are metalloen-zymes consisting of three metal ions, typically two Zn2+ and oneMg2+ ions in their active site. Mammalian APs (mAPs) have low se-quence identity with their bacterial counterparts (25–30%) but theactive site residues and the residues coordinating the metal ions

well studied enzymes both biochemically and structurally (Kimand Wyckoff, 1991). All APs have a basic aba fold and the biolog-ically active species is dimeric. Among the eukaryotic AP homologs,structural information is available for human placental AP (hPLAP)(Le Du et al., 2001; Llinas et al., 2005) and shrimp AP (SAP) (DeBacker et al., 2002, 2004). The overall catalytic mechanism is wellconserved between species. One Zn2+ ion (M1) activates the cata-lytic serine leading to the formation of phosphoserine intermedi-ate. A water molecule activated by the second Zn2+ ion (M2)hydrolyzes the phosphoserine intermediate. The phosphate thengets released or transferred to an acceptor (Xu and Kantrowitz,1991). The third metal site (M3) was originally thought to beresponsible for providing the catalytic base (hydroxide ion boundto Mg2+) which deprotonates the catalytic Ser (Kim and Wyckoff,1991). But recent studies implicate it in stabilizing the transferredphosphate moiety in the transition state (Zalatan et al., 2008).

The differences between the mammalian and prokaryotic APshave been well documented (Zhang et al., 2005). Mammalian APsare more active and less thermostable in comparison to the EcAP.

K. Ghosh et al. / Journal of Structural Biology 184 (2013) 182–192 183

Structurally, the major difference is the presence of a distinctcrown domain and an N-terminal helix swap in the mAPs (Le Duet al., 2001; Bossi et al., 1993). Additionally, mAPs are allostericenzymes (Hoylaerts et al., 1997) and are susceptible to non-competitive inhibition by L-amino acids like, L-Phe, L-Trp, L-Leu,L-homoarginine and levamisole (Fishman and Sie, 1971; Van Belle,1976). There are four major types of APs found in human – germcell (GCAP), intestinal (IAP), placental (PLAP) and tissue non-spe-cific (TNAP). The first three of these are located on chromosome2 and have 90–98% homology in sequence. TNAP is located onchromosome 1 and has �50% homology with the other three iso-forms and is believed to be the ancestor of the tissue-specificAPs. Mutations in TNAP gene has been implicated in hypophospha-tasia involving diseases like, rickets and osteomalacia (Weiss et al.,1988; Henthorn et al., 1992; Mornet, 2000). PLAP, on the otherhand, is thought to be involved in the transfer of maternal IgG tothe fetus (Makiya and Stigbrand, 1992) and also is one of the firstproteins to be ectopically expressed by cancer cells. IAPs have thehighest specific activity among the different isoforms and theyhave been extensively used in diagnosis, immunoassays andmolecular biology (Jablonski et al., 1986; Sekiguchi et al., 2011).

In the drug discovery paradigm, in many instances, poor solubil-ity of the lead molecule precludes its development into a drug.Phosphate prodrugs have been used successfully in few such casesto increase the solubility as well as bioavailability of the parentdrug (Fleisher et al., 1996). IAPs present in the intestinal brush bor-der play an important role in drug metabolism by cleaving phos-phate prodrugs and releasing the parent drug. This approachworks best for the so called class II compounds in biopharmaceuti-cal classification, which have good permeability but poor solubility.However, not all phosphate prodrugs undergo the desired rapidbioconversion by the IAP and there is a need to better understandthis process in vitro to facilitate the design of phosphate prodrugs.Rat and cynomolgus monkey are two of the most common pre-clin-ical species in drug discovery and recently rat intestinal mucosalscraps and Caco-2 monolayers have been used to characterize thephosphate prodrugs in vitro (Haodan et al., 2009). A better andmore quantitative model will be to use pure IAP enzymes. Sincethese are not commercially available, we have cloned, expressedand purified IAPs from human, rat and cynomolgus monkey.

While most species studied have one isoform of IAP, rat is un-ique in having two isoforms of IAP (I and II) (Xie and Alpers,2000), whereas cow has four isoforms of IAP (Manes et al., 1998).Previous work with the rat IAP suggested that the two isoformsare different in terms of their metal ion requirement and showeddifferent inhibitory potential when titrated with high concentra-tion of Zn2+ (Calhau et al., 2000). Jejunal mucosa expressing onlyisoform I was not inhibited by 1 mM Zn2+ whereas the duodenalmucosa expressing both the isoforms was inhibited by 41% in thepresence of 1 mM Zn2+ (Calhau et al., 2000; Harada et al., 2005).

Here, we focus on the biochemical and biophysical character-ization of purified rat and cynomolgus monkey IAP and reportthe first crystal structure of rat IAP. Structural analysis provide arationale for the decreased activity of mammalian APs when ex-pressed in bacteria and further identify a key residue involved inmaking rIAP insensitive to Zn2+. We have also established anin vitro–in vivo correlation of phosphate prodrug cleavage in ratdiscussed in a separate manuscript (Subramanian et al., 2013).

2. Results and discussion

2.1. Expression, purification and characterization of rat andcynomolgus monkey IAP

The rat and cynomolgus monkey IAP genes were cloned as de-tailed in the methods. For expression in insect cells, the original

N-terminal signal sequence was kept intact and the variable C-terminus region (483-end) was replaced by a hexahistidine tag tofacilitate affinity purification of these proteins. The residue num-bering throughout the manuscript is based on the mature proteinsequence. For bacterial expression, the construct was similarto the one used for expression in insect cells except that theN-terminal signal sequence was removed.

APs were purified using metal affinity and size exclusion chro-matography as described in the methods. The purified protein wasa dimer based on the migration in SEC and as determined by thedynamic light scattering (DLS). The purified insect cell expressedrIAP protein was found to be heavily glycosylated and migratedas two distinct bands on SDS–PAGE (Fig. 1a). Both bands were po-sitive in anti-His Western blots indicating them to be the targetprotein (Fig. 1b). We have confirmed their identity by performingan in-gel activity staining. The slower migrating band retainedactivity in the presence of 1% SDS in the gel (Fig. 1c). Since mam-malian APs are known to be glycosylated and dimeric, this suggeststhat the slower migrating band corresponds to a dimer, which isresistant to denaturation. When treated with deglycosylating en-zyme PNGaseF under denaturing conditions, we obtained a singlefaster migrating species on the gel indicating the deglycosylatedmonomers (lane 2, Fig. 1d). The bacterially expressed rIAP proteinwas positive in anti-His Western blots and behaved as dimer in SEC(data not shown).

2.2. Activity of the proteins and effect of metal ions on the activity

Activity of purified APs were tested using p-nitrophenylphos-phate (pNPP) as substrate as described in the methods. The pHoptimum for rIAP and cIAP activity was found to be pH 9.5 (Supple-mentary Fig. S1). Specific activity of the rat, cyno, bovine IAPs andhuman placental AP enzymes tested followed the order bIA-P > rIAP–cIAP > hPLAP (Table 1a). The effect of pH on the catalyticactivity of these enzymes is evident from a 10-fold reduction inthe kcat/Km with an increase in pH from 9.5 to 10.5 (Table 1b).The difference in the rate constant comes from a 10-fold reductionin the Km of the enzymatic reaction at pH 10.5.

The effect of exogenous metal ion on enzyme activity was stud-ied using zinc acetate and magnesium acetate. The enzyme was di-luted in 1 M DEA buffer, pH 9.5 and magnesium acetate wastitrated from 10 lM to 10 mM final concentration. No effect ofexogenous magnesium (up to 10 mM) was observed on rIAP andcIAP activity. Zinc acetate titration at a concentration range of50 lM to 1 mM was performed in 1 M DEA buffer pH 9.5 usingpNPP (10 mM final) as a substrate at 37 �C. Zinc had no effect onthe activity of rIAP but had a pronounced inhibitory effect on cIAP(Fig. 2). Activity of bIAP and hPLAP was also inhibited with increas-ing concentration of Zn2+ (Supplementary Fig. S2). On the otherhand, excess Mg2+ (up to 10 mM) had no effect on the activity ofany of these enzymes under the reaction condition tested (datanot shown).

The inhibitory effect of Zn2+ on APs has been well documentedin the literature (Cathala and Brunel, 1975; Linden et al., 1977). It isbelieved that at higher concentrations, Zn2+ displaces the Mg2+ ionfrom M3 site causing a significant reduction in the activity of theenzyme (Hung and Chang, 2001). Similar to the Zn-bound SAPstructure (De Backer et al., 2002, 2004), rIAP isoform I is hypothe-sized to contain Zn2+ in all 3 metal positions (Harada et al., 2005),accounting for the lack of inhibitory effect when titrated with ex-cess Zn2+. An alternative hypothesis is that rIAP can accommodateeither Zn2+ or Mg2+ in M3 position without much difference inactivity. To understand the metal binding in detail we determinedthe crystal structures of rIAP isoform I expressed in both insect andE. coli cells.

Fig.1. Purification and initial characterization of rIAP: (a) SDS–PAGE analysis of the purified rIAP with coomassie blue staining; 3 and 6 lg of final rIAP protein were loaded inlanes 1 and 2 respectively; molecular weight marker was loaded in lane 3. (b) Anti-His Western blot to confirm the identity of the rIAP protein; both the bands were positiveindicating that may be they are from the same protein. (c) Gel on the left was treated with NBT–BCIP reagent to check for AP activity. The blue band indicates active AP. Theprotein was treated with PNGaseF with or without heat denaturation as noted in the picture. The protein (middle lane) lost activity when heat denatured and treated withPNGaseF and (d) same gel post stained with coomassie blue reagent confirms presence of two different species. (For interpretation of the references to color in this figurelegend, the reader is referred to the web version of this article.)

Table 1aSpecific activity of the different AP enzymes in DEA buffer.

pH 9.5 (IU/mg) pH 10.5 (IU/mg)

Rat IAP-I 254.06 82.04Rat IAP-II 186.83 30.06Rat IAP-I Q319H 226.71 190.29Cyno IAP H319Q 95.85 20.03Cyno IAP 259.71 130.15Human PLAP 0.39 0.43Bovine IAP 652.87 330.47Rat IAP-I E. coli 17.83 –

Fig.2. Effect of exogenous addition of Zn acetate on the activities of rIAP and cIAP:the activity of rIAP and cIAP in the presence of increasing concentration of Zn-acetate was measured by absorbance at 405 nm after 10 min of pNPP hydrolysis.

184 K. Ghosh et al. / Journal of Structural Biology 184 (2013) 182–192

2.3. Structure of rIAP

The rIAP-Ec and rIAP-Ic structures were determined to 2.21 and2.38 Å resolution respectively (Table 2 and Fig. 3). To the best ofour knowledge these are the first crystal structures of any intesti-nal AP.rIAP-Ec structure: In the rIAP-Ec structure, the overall aba APfold, N-terminal helix swap and the disulfide linkages were similarto that of hPLAP (PDB id 1EW2). However, the crown domain(F362-T431) was not visible in the electron density maps(Fig. 3a). The tight interaction between the two monomers keepsthe C-terminal end of the protein together, which is clearly visiblein the structure (colored purple and red in Fig. 3a). Two disulfidelinkages were visible in each monomer between residues C121–C183 and between C467–C474. Only one metal in the active siteat M3 was visible and identified as Mg2+ together with anotherMg2+ (M4) at the metal binding domain (Le Du et al., 2001)(Fig. 3a). The final model includes 2 protein chains, 4 magnesiumions, 2 glycerols and 279 water molecules. Interestingly the active

Table 1bEnzymatic characterization of different IAP enzymes.

Enzyme Km (mM) Vmax (mM/s) kcat (s�1) kcat/Km (M�1s�1)

pH 9.5Rat IAP 1.33 4.35E � 04 1.67E + 03 1.27E + 06Cyno IAP 0.40 2.70E � 04 1.77E + 03 4.46E + 06

pH 10.5Rat IAP 11.896 4.15E � 04 1.60E + 03 1.34E + 05Cyno IAP 3.955 2.26E � 04 1.48E + 03 3.75E + 05Bovine IAP 7.87 7.55E � 05 3.43E + 03 4.36E + 05Human PLAP 1.91 8.49E � 05 1.98E + 01 1.04E + 04

site Ser (92) was covalently linked with phosphate. This could be acrystallization artifact, potentially due to impurities from the re-agents, as phosphate was not present in the crystallization mixturenor was rIAP-Ec crystallized in the presence of pNPP. Moreover, thecrystals were obtained at pH 6.5 which lowers the cleavage rate ofthe phosphoserine bond. In addition, the absence of metal at M2could also account for the trapping of the phosphoserine interme-diate.rIAP-Ic structure: In the rIAP-Ic structure, the overall aba APfold, N-terminal helix swap and the disulfide linkages were similarto that of rIAP-Ec and hPLAP (PDB code 1EW2). The rIAP-Ic proteinwas co-crystallized with pNPP which resulted in a fully metalatedstructure very similar to that of the hPLAP (Le Du et al., 2001)(Fig. 3b and Supplementary Fig. S3). The final model of rIAP-Ic in-cludes 2 protein chains, 4 zinc ions, 6 magnesium ions, 2 pNP (p-ni-tro phenol) molecules, 9 Nag (N-acetyl glucosamine) sugarmolecules and 390 water molecules. We could model all the resi-dues starting from V1 to E482 in the mature protein chains andpart of the C-terminal His-tag as well (4 His residues in monomerA and two His residues in monomer B). The crown domain (F362–T431) was clearly observed in the electron density maps and wasmodeled using the sequence information. Two disulfide linkagessimilar to the rIAP-Ec structure (C121–C183 and C467–C474) wereobserved (Supplementary Fig. S3).

Glycosylation for rIAP-Ic was observed in the electron densitymaps close to N281 and N408 in both the monomers. There are 4Nag units attached to the N281 residue in monomer A and 3

Table 2Data collection and refinement statistics for the two rIAP structures.

Structure refinement

rIAP-Ec rIAP-IcUnit cell (Å) a = 63.32, b = 75.73, c = 170.71 a = 90.31, b = 167.17, c = 71.84Wave length (Å) 1.22 0.9796Resolution (Å) 2.21 (2.28–2.20)a 2.38 (2.47–2.38)Spacegroup P 21 21 21 P 21 21 21

Total No. of reflections measured 251,307 247,490No. of unique reflections 42,064 44,083No. of free reflections 2109 2224Completeness 99.9% 99.9%I/Ir 17.2 (2.4) 10.1 (2.7)R 16.74 16.43Rfree 20.34 20.43No. of protein atoms 6172 7343No. of hetero atoms 313 546No. of solvent atoms 277 390R.m.s. deviation of bond lengths (Å) 0.010 0.010R.m.s. deviation of bond angles (�) 1.08 1.11R.m.s. deviation of dihedral angles (�) 16.74 16.57Wilson B factor (Å2) 42.31 30.20

a Values in the parenthesis are for the last shell.

Fig.3. (a) Structure of rIAP-Ec at 2.21 Å resolution. Monomer A and B are colored as green and yellow respectively. The missing crown domain is circled and indicated by thedotted lines and the rest of the C-terminus is indicated by purple (A) and red (B) respectively. The two disulfide linkages in each monomer are marked by red circles. Mg2+ ionsare shown as green spheres and (b) structure of the rIAP-Ic determined at 2.38 Å resolution. Monomer A and B are colored as green and yellow respectively. Metal ions arerepresented by spheres (Zn – blue, Mg – green), pNP molecule and sugar moieties by sticks.

K. Ghosh et al. / Journal of Structural Biology 184 (2013) 182–192 185

attached to N281 in monomer B. One Nag unit was found attachedto N408 in the crown domain in each of the monomers. The glyco-sylation at N408 is unique and observed for the first time in thisrIAP-Ic structure and, as shown below, might have importantimplication in stability of the dimeric protein. In the hPLAP struc-ture, residues N122 and N249 were glycosylated (Le Du et al.,2001). N122 is conserved between hPLAP and rIAP (SupplementaryFig. S4) and is located on the surface of the protein. Though glyco-sylation is not observed in the electron density maps at N122 forrIAP-Ic, its position mimics that of hPLAP and is most likely glycos-ylated in vivo. Furthermore, while the residue corresponding toN249 of hPLAP is absent in rIAP (Supplementary Fig. S4), glycosyl-ation of N281 could stabilize the additional metal binding site.

2.4. Active site architecture

The active site of each monomer of rIAP-Ic contained 2 Zn2+

atoms (occupancy of 1 and 0.75 for M1 and M2 respectively) and1 Mg2+ ion (M3) along with pNP (Fig. 3b and SupplementaryFig. S5). The protein was crystallized in presence of �50-fold molarexcess of pNPP. This might have facilitated the binding of pNP near

the active site after its generation from the cleavage reaction. TheO5 atom of the nitro group is at a distance of 2.20 Å from the Zn2+

ion at M1. The re-refined structure of hPLAP in presence of p-nitrophenyl phosphonate (pNPPate), a non-hydrolysable substrateanalogue (PDB id 3MK1) also had a pNP molecule bound nearthe active site (Llinas et al., 2005; Stec et al., 2010). Unlike there-refined hPLAP structure or the rIAP-Ec structure, no phosphatedensities were observed in the active site of rIAP-Ic structure.

The M3 site of rIAP-Ic structure was found to be occupied byMg2+ in contrast to a published structural model of rIAP whereM3 was modeled as a Zn2+ (Harada et al., 2005). There was noexogenous Zn2+ in the crystallization trials (a common additivefor most AP crystallization conditions) or during expression andpurification but the purification buffers contained 1 mM MgCl2. Itis possible that continuous exposure to MgCl2 led to the replace-ment of the Zn2+ at M3 with Mg2+ ion. There are structures ofSAP where M3 is either Zn2+ (PDB id 1SHN) or Mg2+ (PDB id1SHQ) and in the SAP/Zn2+structure, H149 (H153 in rIAP) coordi-nates Zn2+ at M3 (De Backer et al., 2002). In the Mg2+ bound SAPstructure (PDB id 1SHQ) De Backer et al. (2004) as well asthe rIAP-Ic structure, H149 in SAP and H153 in rIAP) are

186 K. Ghosh et al. / Journal of Structural Biology 184 (2013) 182–192

superimposable (Fig. 4) and they are not in direct contact with M3but coordinate the Mg2+ ion through water molecules. It was alsoobserved that the R162 side chain in the SAP/Zn2+ structure(1SHN) is in a docked conformation by forming H-bondinginteractions with the bound phosphate (Fig. 4, red dotted lines)(De Backer et al., 2004). On the other hand, in the SAP/Mg2+ struc-ture (1SHQ), the R162 side chain is in a non-docked conformation.The orientation of R166 side chain (in rIAP-Ic) is similar to the ori-entation of the docked R162 side chain in the SAP/Zn2+ structure(1SHN), even though there are no phosphate in the active site, pos-sibly because of ionic interaction between NH1 of R166 and the O6of the pNP (Fig. 4, blue dotted lines). But the H153 side chain inrIAP is in a position similar to the SAP structure with Mg2+ at M3(1SHQ) further indicating the presence of Mg2+ at M3 in ourrIAP-Ic structure (Fig. 4).

Two additional Mg2+ ions were observed in the rIAP-Ic struc-ture. One of these Mg2+ (M4) ions (located�10 Å away from the ac-tive site) was also observed in the rIAP-Ec structure and in thehPLAP structure (PDB id 1EW2) in a site called the metal bindingdomain (Le Du et al., 2001). Coordination of the Mg2+ at this siteby residues E216, E270 and D285 are similar between the rIAPand hPLAP structures. As mentioned in the case of the hPLAP struc-ture (Le Du et al., 2001), it was not possible to distinguish betweenMg2+ and Ca2+ at this position using the current data. However,since excess Mg2+ was utilized during purification and crystalliza-tion, it is most likely that the metal in that position is Mg2+. The 5thMg2+ ion (M5) is unique in the rIAP-Ic structure and was notobserved in the hPLAP structure. It is located near the base of theprotein, at the end of the a11 helix (Supplementary Fig. S6), withan average B-factor of 35 Å2 and is coordinated by three watermolecules and backbone carbonyl atoms of S342 and T345. ThisMg2+ ion is at a distance of >30 Å from the active site and probablyplays a structural role.

2.5. Structural basis of the observed effect of Zn2+ on rIAP

As mentioned earlier, exogenous addition of Zn and Mg acetatehave a different effect on the rIAP in comparison to bIAP, cIAP andhPLAP. Activities of these three IAPs are inhibited in the presence ofexcess Zn2+ whereas there is no effect on rIAP (Fig. 2). The differen-tial effect of excess Zn2+ can be explained by the structural analy-sis. The residues contacting the 3 metal ions in the active sites ofthe AP enzymes have been conserved from bacteria to human(Supplementary Fig. S4). However, there is less conservation in

Fig.4. Comparison of SAP and rIAP active sites: the SAP structures 1SHN (orangecarbon) and 1SHQ (blue carbon) were superimposed onto the rIAP-Ic structure(green carbon); His149 moves towards the Zn at M3 (1SHN) to have a favorableinteraction. R162 has two different orientations (docked and non-docked) and R166in the rIAP structure matches that of the docked conformation (1SHQ). Metal ionsfor 1SHQ are omitted from the picture for simplicity.

the second shell of residues surrounding the metal ions. The M1(Zn2+) site has the highest affinity for the metal and it is mainly be-cause of the coordination through two conserved His residues. TheM2 site has been shown to be less stringent at least in case of EcAP;it has been shown that the M2 Zn2+ can be replaced by Mg2+ with-out affecting the overall activity of the enzyme (kcat/Km) (Tibbittset al., 1996). There is a decrease in affinity for substrate but con-comitantly there is an increase in kcat due to ease of phosphate re-lease in the presence of Zn2+ in M2 (Tibbitts et al., 1996). The M3metal ion, in comparison, is a bit more accessible and, presumably,is the first metal ion to be displaced during titration (Coleman,1992).

Comparison of the structure of hPLAP with rIAP revealed that allthe residues that are in direct contact of the metal ions are con-served and the only difference is in the second sphere of residues(Fig. 5a), where residue 317 was different in rat compared to otherIAPs and hPLAP (Supplementary Fig. S4, position indicated by astar). This was also observed by Harada et al. while discussing astructural model of rIAP (Harada et al., 2005). The side chain ofQ317 is at a distance of 6.64 Å from M3 Mg2+ ion and occupies sim-ilar position as that of H317 in hPLAP (Fig. 5a). In order to confirmthe role of residue 317, we generated the Q317H mutation in rIAPand H317Q mutation in cIAP. They showed a reversal of inhibitoryeffect of Zn2+, i.e. the Q317H mutant rIAP was strongly inhibited byexcess Zn2+ and the H317Q cIAP was resistant to inhibition byZn2+ (Fig. 5b). This indicates that the Q317 residue has a directrole in determining the effect of exogenous Zn2+ on the activityof rIAP.

2.6. Comparison of the two rIAP structures

Based on the superposition of the two rIAP structures (rIAP-Ecand rIAP-Ic) (Supplementary Fig. S6), the TLR tripeptide fragment(residues 368–370) was modeled into the additional 2Fo–Fc densityin the rIAP-Ec structure near the missing crown domain (Supple-mentary Fig. S7). This tripeptide is visible in both the subunits ofthe rIAP-Ec structure and is not connected to the rest of the proteinin the structure. The tripeptide is held in its place by a network ofionic interactions and the tripeptide from monomer B is presentclose to monomer A and vice versa (Fig. 6). The NH1 and NH2 ofthe R370 side chain of monomer B is stabilized by interactions withthe OE1 of E6 of monomer B and the backbone of V85 from mono-mer A. The NE of R370 interacts with E66 OE1 of monomer B. Thebackbone of R370 was stabilized by interaction with the backboneof D86 from monomer A. Calculation of omit map showed cleardensities for the tripeptide (Fig. 6b and c). Overall, the structuresof rIAP-Ec and rIAP-Ic are very similar. Excluding the crown do-main, the r.m.s.d. in Ca main chain atoms between the two struc-tures is 0.58 Å. The number of residues in the dimer interface forthe rIAP-Ic structure is 129 and 80 for the rIAP-Ec structure. The to-tal buried surface areas in the rIAP-Ec and rIAP-Ic structures are2820 (16.7% of total accessible surface area) and 4475 Å2 (23.1%of total accessible surface area) respectively. The difference in thenumber of residues at the interface and consequently the amountof buried surface area stems from the lack of the crown domain inthe rIAP-Ec structure.

In the rIAP-Ic structure, H320, H432 and D316 coordinate theM1 Zn2+ ion while the M2 Zn2+ is stabilized by interactions withD42 and D316. The crown domain is further stabilized by the pres-ence of the Y367 side chain from the other monomer. Additionally,there are two ionic interactions between the backbone of theresidues present in the loop between helices a11 and a12(A321–A324) and residues from the crown domain which stabilizethe region. The side chain NZ of K420 interacts with carbonyl ofA321 and the NE2 of Q421 makes polar interaction with thebackbone carbonyl of G322 (Fig. 7). The Mg2+ at M3 is stabilized

Fig.5. Residue 317: (a) superposition of rIAP (in green) and the hPLAP (in yellow) structures showing the active site. All the residues near the metal ions are conserved exceptfor the change in residue 317 which is about 6.6 Å away from M3 and (b) effect of addition of Zn acetate on the activity of rat Q317H and cynomolgus monkey H317Q IAPmutants.

Fig.6. TLR tripeptide: (a) stabilizing interaction of R370 (magenta) of monomer B (yellow) with neighbouring residues of both monomers A (green) and B are shown by reddotted lines. The distances between the interacting atoms and the residues are labelled. The omit map densities for the TLR tripeptide of monomers A and B at Ir is shown in(b) and (c) respectively.

K. Ghosh et al. / Journal of Structural Biology 184 (2013) 182–192 187

by interaction with D42, S155 and E311 side chains together withwater molecules. On the contrary, in the rIAP-Ec structure, H320and H432 are disordered (Fig. 7) and D316 is moved away(�2.84 Å distance between CG atoms of D316 between the twostructures). Further, owing to the absence of crown domain, theanchoring interaction mediated through residues in the a11 anda12 helices are lost and the loop is disordered allowing the a11 he-lix to move (�10�). This resulted in the displacement of the sidechains of D316 and H320 leading to loss of metal ion at the M2 site.The M3 metal (Mg2+) binding site is intact as the metal coordinat-ing residues are not affected by the loss of the crown domain.

2.7. Why is the crown domain missing from the rIAP-Ec structure?

The purified rIAP-Ec protein was intact and its mass was con-firmed by SDS–PAGE and mass spectrometry (MS) analysis. Theinitial hypothesis for the missing crown domain was that the do-main is floppy in nature and hence it was not visible in the electrondensity maps. Another hypothesis to explain the missing crowndomain was that protein was being cleaved during storage andor crystallization. To explore protein cleavage, crystals were runon SDS–PAGE gels after extensive washing in stabilizing buffer.

We found that the protein migrated faster (at �42kDa) indicatingcleavage during crystallization trials at 20 �C (Fig. 8a). The cleavagewas confirmed by incubating purified protein at room temperaturefor 60 h (Fig. 8b, lane 1) and analyzing samples by mass spec. Themass of the major species was determined to be 42062.7 Da. Wehave also demonstrated that addition of protease inhibitors (Sig-ma) prevents the cleavage. Crystallization attempts in the presenceof the protease inhibitors were unsuccessful indicating cleavage ofrIAP-Ec protein is important in crystallization. This led us tohypothesize that the absence of glycosylation in bacterial expres-sion systems for rIAP may lead to improperly folded crown domainand that this domain, consequently, is more prone to proteasecleavage. As mentioned earlier, there is a T-L-R tripeptide (368–370) fragment from the crown domain visible in the rIAP-Ec struc-ture (Fig. 6). Analysis of the sequence revealed that, cleavage atK388 will produce a fragment with a molecular weight of42,031 Da, which is very close to that observed after MS analysisof the cleaved protein. Under non-denaturing conditions, this firstcleavage is followed by a second cleavage after the R370 (part ofTLR) which becomes exposed leading to the removal of the crowndomain. Chakraborty et al. (2011) recently used CLASP (CataLyticActive Site Prediction) software to show that APs, at least, SAP

Fig.7. Active site comparison between rIAP-Ic and rIAP-Ec structures: superpositionof the rIAP-Ec (blue carbon) and rIAP-Ic (green carbon) structures showing themovement of the a11 helix near the active site of monomer A. The crown domainsare colored in green (A) and yellow (B) respectively for the two monomers of rIAP-Ic. The loop connecting a11 and a12 helices is disordered in rIAP-Ec structure. Inthe rIAP-Ic this loop is stabilized by hydrogen bonding interactions between thecrown domain residues K420 and Q421 and the backbone carbonyl of A321 andG322 respectively. Residues directly contacting M1 (Zn), H320, D316 and H432 sidechains are displaced for the rIAP-Ec structure (shown in blue carbon) as well as D42which contacts both M2 and M3. The Y367 present close to the TLR peptide frommonomer B (yellow) is also shown in the rIAP-Ic structure.

188 K. Ghosh et al. / Journal of Structural Biology 184 (2013) 182–192

has partial proteolytic activity where it can cleave self as well asother proteins. Moreover, its autocatalytic activity increases whenthe metal ions are removed using EDTA. This indicates that therIAP-Ec in absence of the fully folded crown domain can potentiallyauto-cleave to release the crown domain.

Fig.8. Why crown domain is missing from rIAP-Ec structure? (a) one of the rIAP-Eccrystals is loaded onto a SDS–PAGE (lane 1) after washing it thoroughly in reservoirbuffer. It migrated corresponding to molecular weight of �40 kDa protein distinctlyfaster than the original purified protein (lane 2) which migrates as �55 kDa and (b)result of incubating the rIAP-Ec protein with and without protease inhibitors for66 h. Lane 1 – rIAP incubated at 20 �C without PI; lane 2 – rIAP with PI incubated at20 �C; lane 3 – rIAP without PI incubated at 4 �C. The faster migrating band in lane 1was later confirmed by ESI-MS analysis as 42062.7 Da.

On the other hand, the K388 residue in the rIAP-Ic structure isshielded by the glycosylation at the N408 residue in the other sub-unit. It could be the reason why the crown domain is protectedwhen expressed in Sf9 cells. Further analysis of the rIAP protein se-quence revealed that there are three possible N-linked glycosyla-tion motifs (N-X-S/T) at residues N122, N281 and N408respectively (Supplementary Fig. S4). To test this hypothesis, theseasparagine residues were mutated individually to aspartic acid anda triple mutant was also generated (N122D/N281D/N408D). Theywere expressed in Sf9 cells and purified from media. Stability ofthe purified proteins was monitored using a fluorescence basedthermal shift assay (TSA) (Pantoliano et al., 2001; Lo et al., 2004).The results are summarized in Table 3. The Tm of IAPs purified frominsect cells were dependent on the buffer conditions and, in mostcases, the proteins were less stable in DEA buffer (di-ethyl amine)pH 9.5 and were stabilized with the addition of Zn2+. The proteinswere most stable in the storage buffer (100 mM Tris (pH 7.4),300 mM NaCl, 1 mM MgCl2 and 10% Glycerol (v/v)). Interestingly,the rIAP-Ec was found to be significantly less stable specificallyin the DEA buffer pH 9.5 (Tm = 37 �C). The Tm of the N to D mutantsdemonstrated reduced thermal stability in comparison to the wildtype rIAP (Table 3) and thermal stability profile matches exactly tothe rIAP expressed in E. coli. The reduction in thermal stabilitycould possibly be due to the lack of N-linked glycosylation. Theactivity of the triple mutant is reduced to half of that observedfor the wild type (data not shown). The activity measurementswere performed at 37 �C and the Tm of the protein is also 37 �C.This could possibly be explained by two hypotheses – one wherethe insect cell is better equipped to fold the crown domain evenin the absence of glycosylation or the glycosylation is not essentialfor proper folding of the crown domain. Based on our results webelieve that glycosylation plays an important role in the properfolding of the crown domain.

We have isolated the rIAP-Ec protein without the crown domain(Fig. 8b) and compared its activity with that of the intact rIAP-Ec(either at 4 �C or incubated with protease inhibitor cocktail at roomtemperature). The specific activity of the intact rIAP-Ec protein issimilar to that of the cleaved product in presence of 1 mM Zn ace-tate (�16 IU/mg). The protease inhibitor did not alter the activityof the rIAP-Ic protein when tested under similar conditions indicat-ing that the phosphatase activity is not affected by the proteaseinhibitor. This result indicates that the activity of rIAP-Ec proteinis unaffected by the presence or absence of the crown domain,leading us to believe that the crown domain is not folded properlyin the rIAP-Ec protein. Consequently, the active site was notformed properly and the affinity towards the metal ions was de-creased leading to lowering of activity. In presence of excess Zn2+

Table 3Thermal stability of different APs.

Name of Protein Tm DEA pH9.5 (�C)

Tm DEA pH9.5 + 1 mM Zn2+ (�C)

Tm SBa pH7.4 (�C)

hPLAP 71 63 NDb

bIAP 60 59 65.5rIAP-I 60 66.5 65.5rIAP-II 39 62.5 NDrIAP-Ec 37 64 57cIAP 38.5/60.5 62.5 41/65rIAPN281D 42/54.5 66 64.5rIAPN408D 41/54.5 65.5 55.5/63rIAPN281D N408D 39.5/54 65.5 50/62.5rIAP TM (N122D

N281D N408D)37/47 65.5 51/61.5

Two values in a single column for some of the constructs indicate dual transitions.a SB – storage buffer.b ND – not determined.

Fig.9. Protease cleavage sites: superposition of the two rIAP structures showing location of the K386, N408 and R370 residues. The K386 in ball and stick is shielded by theglycosylation (sticks) at N408. Cleavage after K386 will lead to a fragment with theoretical molecular weight of 42,031 Da.

K. Ghosh et al. / Journal of Structural Biology 184 (2013) 182–192 189

the active site could be partially occupied by Zn2+ leading to partialrecovery of activity (4-fold for intact rIAP-Ec and 30-fold for rIAP-Ec without the crown domain, data not shown).

3. Conclusion

Rat and cynomolgus monkey IAPs were cloned, expressed, puri-fied and characterized. These enzymes are being used to test thecleavage efficiency of phosphate prodrugs. The generated data willhelp us understand the difference in cleavage efficiency betweenspecies. The crystal structure of rIAP was determined and, to thebest of our knowledge, this is the first report of expression andcrystallization of recombinant rIAP. The crown domain in theE. coli expressed rIAP protein is not properly folded and is proneto protease cleavage. Based on recent literature reports, the lossof crown domain due to protease cleavage could result from self-cleavage of the protein, specifically when metal sites are not fullyoccupied (Chakraborty et al., 2011). Lack of properly folded crowndomain leads to the movement of key residues involved in interac-tion with the active site metal ions which in turn is responsible forthe decrease in activity. The susceptibility to proteases is decreasedin case of rIAP expressed in Sf9 cells because of N408 glycosylationwhich protects K386 from cleavage (Fig. 9).

Activity of rIAP was not inhibited by excess Zn2+ leading to thehypothesis that rIAP has Zn2+ in all three metal positions. Crystalstructure of rIAP protein contained Mg2+ at the M3 position, butit does not rule out the possibility that Zn2+ can also occupy thisposition without changing the activity of the enzyme. The lack ofinhibition by excess Zn2+ in rIAP was reversed in the Q317H mu-tant. On the other hand, the corresponding H317Q mutant of cIAPwas not inhibited by excess Zn2+ supporting the hypothesis thatthe glutamine residue at position 317 renders the rIAP insensitiveto Zn2+ probably by indirect interaction mediated by water mole-cules. The structural and biochemical data together suggests twopossibilities – one indicating rIAP might be able to accommodateeither Zn2+ or Mg2+ in its active site without losing activity andthe other indicating rIAP has reduced affinity for Zn2+ specificallyat the M3 site, hence its activity is not affected by excess Zn2+.

Residue Q317 in rIAP which is >6 Å away from the M3 site and�4 Å from H153, plays a role in this process (indicated by ourmutagenesis data) plausibly through one or more water molecules.

4. Materials and methods

Sequence of all the primers used in this study can be found inthe Supplementary document 1.

4.1. Denovo cloning

Rat and cynomolgus monkey total intestinal RNA was pur-chased from Biochain. Rat IAP sequence from gene bank(NM_022665) was used to design primers for cloning. For cyno-molgus monkey, primers were designed in conserved regions usingmultiple sequence alignment of gorilla, rhesus, human and chimpsequences (all primers used in the study are listed in Supplemen-tary document 1). First strand cDNA was synthesised using Super-script III cDNA synthesis kit (Invitrogen). Independent amplicons ofexpected size (�1.7 kb) obtained from PCR amplification wascloned into pTZ57R/T vector (InsTAcloning kit, Thermo Scientific)and sequence confirmed (pTZ57RT/AP). For expression in insectcells, the wild type IAP construct (up to residue 482 of mature se-quence, Supplementary Fig. S4) was subcloned into pFastBac (pFB)vector (Invitrogen). An NdeI site was created just upstream of theATG start codon for cloning purposes in the 50-primer and the30-primer had an XhoI site and the His6 tag and the IAP gene wasPCR amplified using pTZ57RT/AP as template. The resultant NdeIand XhoI fragment was inserted into the NdeI and XhoI sites ofthe pFastBac expression vector. The recombinant bacmid DNAwere transfected into Sf9 cells. After two rounds of amplification,the baculovirus stock was transfected into Sf9 cells for proteinexpression.

4.2. Subcloning for expression in E. coli

Region comprising residues 1–482 of rat IAP without theN-terminal signal sequence was subcloned into pET28 vector for

190 K. Ghosh et al. / Journal of Structural Biology 184 (2013) 182–192

expressing the IAP protein in E. coli Rosetta-gami cell. Six His res-idues were included from the vector in the C-terminus for affinitypurification.

4.3. IAP mutagenesis

The pFastBac clones of rIAP and cIAP were used for mutagenesisusing QuikChange Site Directed Mutagenesis Kit (Agilent) as permanufacturer’s recommended protocol (primers used are listedin Supplementary document 1). Mutations (Rat: Q317H, N301D,N428D, N142D, Cyno: H317Q) were confirmed by DNA sequencing.Double and triple mutants were generated subsequently as re-quired using the single mutants as templates. Bacmids were gener-ated following similar protocol.

4.4. Expression and purification in insect cell

Sf9 cells at 2 � 106 cells/ml was infected at 27 �C for 66 h withP2 virus stock (30 ml/L) of respective IAP constructs. Cells wereharvested by centrifugation at 2500g for 25 min. Secretion of theIAP protein into the media was confirmed by measuring activityusing NBT/BCIP reagents (see below). Supernatant was filteredthrough 0.22 lm bottle top filter(s) and pass onto Q-Sepharosechromatography. The flow through containing the AP activitywas subjected to Ni–nitrilotriacetic acid (Ni–NTA) chromatogra-phy. The Ni–NTA eluted fractions containing AP activity werepooled and subjected to size exclusion chromatography (SEC)using Superdex 200 HR 26/60 column (GE Healthcare) in storagebuffer (100 mM Tris (pH 7.4), 300 mM NaCl, 1 mM MgCl2 and10% Glycerol (v/v)). All the buffers used in purification had 1 mMMgCl2.

4.5. In-gel activity assay

Reaction of AP with the combination of BCIP (5-Bromo-4-Chloro-30-Indolylphosphate p-Toluidine Salt) and NBT (Nitro-BlueTetrazolium Chloride) to yield an intense, insoluble black-purpleprecipitate was used to measure AP activity in-gel. The proteinsamples were ran on Novex 4–20% pre-cast Tris–Glycine gradientgels (Invitrogen). The samples were not heat denatured but had0.5% SDS in the sample loading dye along with 5 mM 2-mercap-toethanol. Gels were run on the recommended Tris–Glycine run-ning buffer containing 1% SDS. Post completion of the run, thegel was incubated in 10 ml 1� PBS containing 100 ll Reagent-Aand 100 ll Reagent B (AP Conjugate Substrate Kit #170-6432, Col-orimetric AP substrate reagent kit, includes premixed BCIP andNBT solutions, Bio-Rad Laboratories) as per manufacturer’s guide-lines. Active proteins were indicated by the appearance of blue col-or bands on the gel after some time. The gel was then washed withwater, scanned and subsequently stained with coomassie brilliantblue staining (Invitrogen).

4.6. Deglycosylation with PNGaseF

Deglycosylation reaction was performed with PNGaseF (P7367Sigma) under native and denatured conditions. 10 lg of respectiveIAP samples were used for each reaction. For denaturation, sampleswere boiled at 100 �C for 5 min in presence of 5% SDS and 0.4 MDTT. 2 ll of 50 mM sodium phosphate buffer pH 7.5 and 2.0 ll of10% NP-40 was added to the sample and mixed followed by addi-tion of 2 ll PNGaseF (1 Unit). Total volume of the reaction wasmade up to 20 ll with water and the mixture was incubated atroom temperature for 3 h. For native conditions, 10 lg of IAP sam-ples was mixed with 50 mM sodium phosphate buffer, pH 7.5 andincubated with 2 Units of PNGaseF at room temperature for 3 h

prior to run on a SDS–PAGE followed by in-gel activity test as men-tioned above.

4.7. Thermal stability analysis

Thermal stability of the different variants of IAP’s was per-formed using Sypro Orange dye (Molecular Devices) and Bio-Radi5 thermal cycler. The protein concentration was constant at0.4 mg/ml in all the measurements and each data point was repli-cated at least twice.

Dynamic light scattering measurements were performed usingDynapro 96-well plate reader from Wyatt Technology Corpora-tions. All measurements were made at protein concentration of1 mg/ml.

Mass spectrometric analysis: Total mass estimation of proteinsamples were performed on a Bruker HCT ultra ESI-MS coupledwith Agilent 1200 series HPLC system.

4.8. Activity measurements

Bovine intestinal alkaline phosphatase (Bovine IAP) and humanplacental alkaline phosphatase (hPLAP) were purchased fromSigma (P6774, P1391 respectively). The specific activity of theenzymes was determined in 1 M DEA (diethanolamine), 5 mMMg-acetate buffer using 10 mM final concentration of p-nitrophe-nylphosphate (pNPP) as a substrate and at respective pH condi-tions at 37 �C. The reaction was monitored at 405 nm using aSpectramax plate reader for the p-nitrophenol product. One unitof alkaline phosphatase activity is defined as the amount ofenzyme that hydrolyzes 1 lmol of pNPP per min at 37 �C at a givenpH. The kinetics of phosphate hydrolysis was measured for rat IAP,cyno IAP, human placental AP and bovine IAP in 1 M DEA, 5 mMMg acetate buffer using pNPP. The effect of pH on enzyme activitywas monitored using 10 mM pNPP (final) in 100 mM citrate (pH6.0), 100 mM Tris–HCl (pH 7.0, 7.4), 100 mM Tris- (pH 8.0, 8.5,9.0), 1 M DEA (pH 9.5, 10, 10.5, 11.0) respectively.

The effect of exogenous metal ion on enzyme activity was stud-ied using zinc acetate and magnesium acetate. 0.0012 IU of eachenzyme in 1 M DEA, pH 9.5 buffer was used to assess the effectof exogenous metal ions (Zn2+ and Mg2+) on activity. Magnesiumacetate (Sigma) was titrated from 10 lM to 10 mM final concentra-tion. Zinc acetate (Sigma) was titrated from 50 lM to 1 mM in 1 MDEA buffer pH 9.5 using pNPP (10 mM final) as a substrate at 37 �C.

4.9. Crystallization of rIAP

Initial crystallization trials were performed with 10 mg/ml pro-tein at 20 �C by sitting-drop vapor diffusion method in 96-wellcrystallization plates with commercially available crystallizationscreens (Hampton Research). Both rIAP-Ec as well as rIAP-Icshowed signs of crystallization in several conditions. The bestrIAP-Ec crystals were obtained with 0.1 M NH4Cl, 0.1 M MES,pH 6.5 and 20% PEG MME 2000 as the reservoir solution. TherIAP-Ic was co-crystallized with 10 mM pNPP using 1.4 M sodiumcitrate, 0.1 M Hepes, pH 7.5 as the reservoir solution. Rod shapedcrystals appeared for rIAP-Ec and needle shaped crystals wereobtained for rIAP-Ic after 4 days. The crystals were soaked in thereservoir solution containing 20% glycerol and flash frozen in liquidnitrogen for data collection.

4.10. Data collection

X-ray diffraction images for rIAP-Ec were obtained at 2.20 Å res-olution using SER-CAT 22ID beam line at Argonne National Labora-tory, Argonne IL, USA. Data for rIAP-Ic crystal was collected usingCLS_CMCF1_08ID beamline of Canadian light source at 2.38 Å

K. Ghosh et al. / Journal of Structural Biology 184 (2013) 182–192 191

resolution. The images were processed using Denzo and Scalepackin the HKL-2000 program package (Otwinowski and Minor, 1997).

4.11. Structure solution and refinement

Conversion of the integrated intensities (I) to the structure fac-tors (F) was done using TRUNCATE and molecular replacement wasdone with PHASER, both from the CCP4 program suite (Collabora-tive Computational Project N, 1994).

4.12. rIAP-Ec

Initial trials of Molecular replacement with hPLAP (1EW2) wereunsuccessful yielding no solution. A new model of hPLAP (1EW2)was made where residues at N-terminal and C-terminal ends weretrimmed down and where the crown domain was removed. Thisyielded a solution with a dimer in the asymmetric unit and solventcontent of 47%. All the model building was done in COOT (Emsleyand Cowtan, 2004). Program Auto Buster (version 1.11.1) (Blancet al., 2004) was used for the positional refinement. Presence ofunmodeled density near the M3 site in the 2Fo–Fc and Fo–Fc mapindicated presence of metal ion. Initially, Zn2+ was modeled intothe density and refined. Later Zn2+ was replaced with Mg2+ ionbased on the B-factors (Mg2+ B-fac �70 Å2, Zn2+ B-fac �130 Å2),electron density and co-ordination. Crown domain (Phe 362–Thr461) was completely absent in the electron density maps, henceleft unmodeled. After several round of the model building/correc-tions and refinement the final Rwork and Rfree of the structure were16.74 and 20.34 respectively.

4.13. rIAP-Ic

Initial trials of molecular replacement with hPLAP (1EW2) usingMOLREP (Vagin and Teplyakov, 1997) yielded a solution with a di-mer in asymmetric unit and solvent content of 50%. Rigid bodyrefinement resulted in Rwork and Rfree of 36 and 36 respectively.The electron density maps were evaluated with the program COOT(Emsley and Cowtan, 2004) and the model was refined with AutoBuster (version 1.11.1) (Blanc et al., 2004). After several round ofthe model building/corrections and refinement the final Rwork

and Rfree of the structure were 16.60 and 20.32 respectively. The fi-nal model includes 2 protein chains, 4 zinc ions, 6 magnesium ions,2 pNP molecules, 9 Nag sugar molecules and 390 water molecules.The data collection and refinement statistics are summarized inTable 2.

4.14. Quality of the models

Program PROCHECK (Laskowski et al., 1993) in the CCP4 pack-age was used to check the geometry of the refined model. TheRamachandran statistics shows 93.3% of residues in core regionand 0.0% of residues in disallowed regions for rIAP-Ec and 90.9%of residues in core region and 0.0% of residues in disallowed re-gions for rIAP-Ic. No D-amino acids or incorrect chiral volumeswere observed in the final models. Models were validated usingMolprobity (Davis et al., 2007). Pictures of the structural modelswere generated using Pymol (DeLano, 2002).

4.15. Protein data bank entry codes

The coordinates have been deposited in the PDB at Brookhavenwith entry codes: rIAP-Ec, 4KJD; rIAP-Ic, 4KJG.

Acknowledgments

The authors would like to thank Shamrock Structures LLC fortheir high quality crystallographic data collection services. Weare thankful to Yoganand Vadari for his help in the initial stagesof the project and Drs. Mike Sinz, Jodi Muckelbauer and SucharitaBose for critical reading of the manuscript. We would also like toacknowledge inputs and advices from Drs. James Bryson, MianGao, Sandhya Mandlekar, Kimberley Lentz and David Rodriguesat different stages of the project.

Appendix A. Supplementary data

Supplementary data associated with this article can be found, inthe online version, at http://dx.doi.org/10.1016/j.jsb.2013.09.017.

References

Blanc, E., Roversi, P., Vonrhein, C., Flensburg, C., Lea, S.M., et al., 2004. Refinement ofseverely incomplete structures with maximum likelihood in BUSTER-TNT. ActaCrystallogr. D: Biol. Crystallogr. 60, 2210–2221.

Bossi, M., Hoylaerts, M.F., Millan, J.L., 1993. Modifications in a flexible surface loopmodulate the isoenzyme-specific properties of mammalian alkalinephosphatase. J. Biol. Chem. 268, 25409–25416.

Calhau, C., Martel, F., Hipolito-Reis, C., Azevedo, I., 2000. Differences betweenduodenal and jejunal rat alkaline phosphatase. Clin. Biochem. 33, 571–577.

Cathala, G., Brunel, C., 1975. Bovine kidney alkaline phosphatase. Catalyticproperties, subunit interactions in the catalytic process, and mechanism ofMg2+ stimulation. J. Biol. Chem. 250, 6046–6053.

Chakraborty, S., Minda, R., Salaye, L., Bhattacharjee, S.K., Rao, B.J., 2011. Active sitedetection by spatial conformity and electrostatic analysis—unravelling aproteolytic function in shrimp alkaline phosphatase. PLoS ONE 6 (12), e28470.

Coleman, J.E., 1992. Structure and mechanism of alkaline phosphatase. Annu. Rev.Biophys. Biomol. Struct. 21, 441–483.

Collaborative Computational Project N, 1994. The CCP4 suite: programs for proteincrystallography. Acta Crystallogr. D 50, 760–763.

Davis, I.W., Leaver-Fay, A., Chen, V.B., Block, J.N., Kapral, G.J., et al., 2007. MolProbity:all-atom contacts and structure validation for proteins and nucleic acids. Nucl.Acids Res. 35, W375–W383.

De Backer, M., McSweeney, S., Rasmussen, H.B., Riise, B.W., Lindley, P., Hough, E.,2002. The 1.9 Å crystal structure of heat-labile shrimp alkaline phosphatase. J.Mol. Biol. 318, 1265–1274.

De Backer, M.M.E., McSweeney, S., Lindley, P.F., Hough, E., 2004. Ligand-binding andmetal exchange crystallographic studies on shrimp alkaline phosphatase. ActaCrystallogr. D 60, 1555–1561.

DeLano, W.L., 2002. The PyMOL Molecular Graphics System. DeLano Scientific, PaloAlto, CA.

Emsley, P., Cowtan, K., 2004. Coot: model-building tools for molecular graphics.Acta Crystallogr. D: Biol. Crystallogr. 58, 2126–2132.

Fishman, W.H., Sie, H.G., 1971. Organ-specific inhibition of human alkalinephosphatase isoenzymes of liver, bone, intestine and placenta; L-phenylalanine, L-tryptophan and L-homoarginine. Enzymologia 41, 140–167.

Fleisher, D., Bong, R., Stewart, B.H., 1996. Improved oral drug delivery: solubilitylimitations overcome by the use of prodrugs. Adv. Drug Deliv. Rev. 19, 115–130.

Haodan, Y., Na, L., Lai, Y., 2009. Evaluation of in vitro models for screening alkalinephosphatase-mediated bioconversion of phosphate ester prodrugs. Drug Metab.Dispos. 37, 1443–1447.

Harada, T., Koyama, I., Matsunaga, T., Kikuno, A., Kasahara, T., Hassimoto, M., Alpers,D.H., Komoda, T., 2005. Characterization of structural and catalytic differencesin rat intestinal alkaline phosphatase isozymes. FEBS J. 272, 2477–2486.

Henthorn, P.S., Raducha, M., Fedde, K., Lafferty, M.A., Whyte, M.P., 1992. Differentmissense mutations at the tissue-nonspecific alkaline phosphatase gene locusin autosomal recessively inherited forms of mild and severe hypophosphatasia.Proc. Natl. Acad. Sci. USA 89, 9924–9928.

Hoylaerts, M.F., Manes, T., Millan, J.L., 1997. Mammalian alkaline phosphatases areallosteric enzymes. J. Biol. Chem. 272, 22781–22787.

Hung, H.C., Chang, G.G., 2001. Differentiation of the slow-binding mechanism formagnesium ion activation and zinc ion inhibition of human placental alkalinephosphatase. Protein Sci. 10, 34–45.

Jablonski, E., Moomaw, E.W., Tullis, R.H., Ruth, J.L., 1986. Preparation ofoligodeoxynucleotide-alkaline phosphatase conjugates and their usehybridization probes. Nucl. Acids Res. 14, 6115–6128.

Kim, E.E., Wyckoff, H.W., 1991. Reaction mechanism of alkaline phosphatase basedon crystal structures. Two-metal ion catalysis. J. Mol. Biol. 218, 449–464.

Laskowski, R.A., Macarthur, M.W., Moss, D.S., Thornton, J.M., 1993. PROCHECK: aprogram to check the stereochemical quality of protein structures. J. Appl.Crystallogr. 26, 283–291.

Le Du, M.H., Stigbrand, T., Taussig, M.J., Menez, A., Stura, E.A., 2001. Crystal structureof alkaline phosphatase from human placenta at 1.8 Å resolution. J. Biol. Chem.276, 9158–9165.

192 K. Ghosh et al. / Journal of Structural Biology 184 (2013) 182–192

Linden, G., Chappelet-Tordo, D., Lazdunski, M., 1977. Milk alkaline phosphatase.Stimulation by Mg2+ and properties of the Mg2+ site. Biochim. Biophys. Acta483, 100–106.

Llinas, P., Stura, E.A., Menez, A., Kiss, Z., Stigbrand, T., Millan, J.L., Le Du, M.H., 2005.Structural studies of human placental alkaline phosphatase in complex withfunctional ligands. J. Mol. Biol. 350, 441–451.

Lo, M.C., Aulabaugh, A., Jin, G.X., Cowling, R., Bard, J., Malamas, M., Ellestad, G., 2004.Evaluation of fluorescence-based thermal shift assays for hit identification indrug discovery. Anal. Biochem. 332, 153–159.

Makiya, R., Stigbrand, T., 1992. Placental alkaline phosphatase has a binding site forthe human immunoglobulin-G FC portion. Eur. J. Biochem. 205, 341–345.

Manes, T., Hoylaerts, M.F., Muller, R., Lottspeich, F., Holke, W., Millan, J.L., 1998.Genetic complexity, structure, and characterization of highly active bovineintestinal alkaline phosphatases. J. Biol. Chem. 273, 23353–23360.

Mornet, E., 2000. Hypophosphatasia: the mutations in the tissue-nonspecificalkaline phosphatase gene. Hum. Mutat. 15, 309–315.

Otwinowski, Z., Minor, W., 1997. Processing of X-ray diffraction data collected inoscillation mode. Method Enzymol. 276, 307–326.

Pantoliano, M.W., Petrella, E.C., Kwasnoski, J.D., Lobanov, V.S., Myslik, J., Graf, E.,Carver, T., Asel, E., Springer, B.A., Lane, P., Salemme, F.R., 2001. High-densityminiaturized thermal shift assays as a general strategy for drug discovery. J.Biomol. Screen. 6, 429–440.

Sekiguchi, S., Kohno, H., Yasukawa, K., Inouye, K., 2011. Chemiluminescent enzymeimmunoassay for measuring leptin. Biosci. Biotechnol. Biochem. 75 (4), 752–756.

Stec, B., Cheltsov, A., Millan, J.L., 2010. Refined structures of placental alkalinephosphatase show a consistent pattern of interactions at the peripheral site.Acta Crystallogr. F66, 866–870.

Subramanian, M., Paruchury, S., Singh, S., Singh, S., Arla, R., Pahwa, S., Jana, S.,Katnapally, P., Vadari, Y., Laksmaiah, B., Mazumder, D., Ghosh, K., Marathe, P.,Mandlekar, S., 2013. Characterization of recombinantly expressed rat andmonkey intestinal alkaline phosphatases: in vitro studies and in vivocorrelations. DMD Fast Forward 1, 1–4.

Tibbitts, T.T., Murphy, J.E., Kantrowitz, E.R., 1996. Kinetic and structuralconsequences of replacing the aspartate bridge by asparagine in the catalyticmetal triad of Escherichia coli alkaline phosphatase. J. Mol. Biol. 257, 700–715.

Vagin, A., Teplyakov, A., 1997. MOLREP: an automated program for molecularreplacement. J. Appl. Crystallogr. vol. 30, 1022–1025.

Van Belle, H., 1976. Alkaline phosphatase. I. Kinetics and inhibition by levamisole ofpurified isoenzymes from humans. Clin. Chem. 22, 972–976.

Weiss, M., Cole, D.E.C., Ray, K., Whyte, M., Laffeerty, M.A., Mulivor, R.A., Harris, H.,1988. A missense mutation in the human liver/bone/kidney alkalinephosphatase gene causing a lethal form of hypophosphatasia. Proc. Natl.Acad. Sci. USA 85, 7666–7669.

Xie, Q., Alpers, D.H., 2000. The two isozymes of rat intestinal alkaline phosphataseare products of two distinct genes. Physiol. Genomics 3, 1–8.

Xu, X., Kantrowitz, E.R., 1991. A water-mediated salt link in the catalytic site ofEscherichia coli alkaline phosphatase may influence activity. Biochemistry 30,7789–7796.

Zalatan, J.G., Fenn, T.D., Herschlag, D., 2008. Comparative enzymology in thealkaline phosphatase superfamily to determine the catalytic role of an activesite metal ion. J. Mol. Biol. 384, 1174–1189.

Zhang, L., Buchet, R., Azzar, G., 2005. Distinct structure and activity recoveries revealdifferences in metal binding between mammalian and Escherichia coli alkalinephosphatases. Biochem. J. 392, 407–441.