Polaina J., MacCabe A.P. (ed.). Industrial Enzymes. Structure, Function and Applications

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PENICILLIN ACYLASES 597

Verhaert, R.D.M., Riemans, A.M., Laan, J., Duin, J. and Quax, W.J. (1997). Molecular cloning and

analysis of the gene encoding the thermostable penicillin G acylase from Alcaligenes faecalis. Appl.

Environ. Microbiol. 63, 3412–3418.

Vohra, P.K., Sharma, R., Kashyap, D.R. and Tewari, R. (2001). Enhanced production of penicillin G

acylase from a recombinant Escherichia coli. Biotechnol. Letts. 23, 531–535.

Wegman, M.A., Janssen, M.H.A., Van Rantwijk, F. and Sheldon R.A. (2001). Towards biocatalytic

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CHAPTER 34

HYDANTOINASES

YUN-PENG CHAO

1∗

, CHUNG-JEN CHIANG

, JONG-TZER CHERN

AND JASON T.C. TZEN

Department of Chemical Engineering, Feng Chia University

Department of Life Science, and

Graduate Institute of Biotechnology, National Chung-Hsing University, Taichung, Taiwan, ROC

∗

ypchao@fcu.edu.tw

1. INTRODUCTION

The hydantoin-cleaving enzymes were first discovered in animals and later charac-

terized using partially purified enzymes from animals and plants (Eadie et al.,

1949; Gaebler and Keltch, 1926). Assigned by the same EC number (EC 3.5.2.2),

hydantoinase (HYD) and dihydropyrimidase (DHP) mediate a similar type of

hydrolytic reaction but exhibit different substrate specificities and kinetic charac-

teristics. They catalyse the reversible hydrolytic ring opening of the amide bond

(-CO-NH-) in five- or six-membered cyclic diamides (Fig. 1). Hydrolysis of the

six-membered ring by DHP is the second step in reductive catabolism of pyrim-

idines in human. This led to the suggestion that HYD may act like the microbial

counterpart (Vogels and Van der Drift, 1976). It was later found that the hydrolysis

of 5-substituted hydantoins by microbial HYDs produced enantiomerically pure

N-carbamoyl amino acids. These intermediates could be further converted to D- or

L-amino acids in an irreversible reaction catalysed by the highly stereoselective

microbial N-carbamoylases (Olivieri et al., 1979). This finding has ushered in a

biotechnological interest in the production of optically pure amino acids because of

their potential for use in the production of antibiotics, peptide hormones, pyrethroids

and pesticides (Owgawa and Shimizu, 2002).

Given the industrial importance of HYDs, natural habitats were intensively

screened for the presence of microbes having high levels of HYD activity. By

the conventional approach of using hydantoin or its derivatives as sole carbon and

nitrogen sources, many microbes with active HYD activity were identified (Yamada

et al., 1978). This screening revealed HYDs to be widely distributed among different

599

J. Polaina and A.P. MacCabe (eds.), Industrial Enzymes, 599–606.

600 YUN-PENG CHAO ET AL.

Figure 1. Typical hydrolysis reactions catalysed by (1) dihydropyrimidase and (2) hydantoinase

genera of bacteria. Microbial HYDs display distinct substrate specificities and are

generally classified depending on their stereoselectivity. However, this classification

is sometimes misleading since it does not provide any information linking the evolu-

tionary relationship of distinct HYDs with their enantioselectivities. For instance,

the HYD from Flavobacterium sp. was found to be L-selective for indolymethyl-

hydantoin but D-selective for benzyloxymethylhydantoin (Yokozeki et al., 1987).

2. BIOLOGICAL FUNCTION AND MOLECULAR STRUCTURE

Whilst a wide variety of HYDs exists in micro-organisms their biological function

remains largely unknown. Nevertheless, with the help of computer analysis the

amino acid sequences of 16 amidohydrolases (AHL) were aligned and compared

(Kim and Kim, 1998). It was found that a long region containing invariant amino

acid residues shared 69% similarity among the enzymes analysed. As a result

of an extensive survey of the protein data bank, a particular sequence consisting

of one aspartic acid and four histidine residues was identified as being highly

conserved in the functionally related AHL. Introduction of mutations into this

conserved region of the D-HYD gene of Bacillus stearothermophilus SD1 (HYDbs)

revealed that these invariant residues play a catalytic role. This has ultimately led

to the grouping of DHP, allantoinase, dihydroorotase (DHO) and HYD into the

HYDANTOINASES 601

cyclic AHL superfamily due to their similar catalytic functions and rigidity in the

conserved sequence (Kim and Kim, 1998).

Our understanding of the catalytic mechanism of HYD has been greatly improved

by the recent availability of the crystal structure of HYDbs (Cheon et al., 2002).

Refined at 3.4 Å resolution, the three-dimensional structure of HYDbs shows that

each subunit assumes a classic TIM barrel fold and an extra domain. Struc-

tural comparison reveals that HYDbs shares a strikingly resemblance to DHO of

Escherichia coli. This structural similarity also extends to the active site, thereby

suggesting that HYDbs is a di-zinc metalloenzyme: one metal ion activates an

attacking hydroxyl while the second metal ion stabilizes the negative charge of the

carbonyl oxygen (Thoden et al., 2001). Furthermore, the substrate binding pocket

of HYDbs was identified by structural suuperimposition of these two enzymes.

From a close-up stereodiagram, three stereochemistry gate loops (SGLs) could be

clearly discerned. These SGLs were proposed to constitute the hydrophobic core

of the substrate binding site and be responsible for the recognition of the exocyclic

substituents of hydantoin. This has been evidenced recently by a study that has

shown that manipulation of the amino acid residues in the SGLs of HYDbs induces

a remarkable change in substrate specificity (Cheon et al., 2004). In particular,

the enzymatic affinity of mutant HYDbs towards the aromatic substrate hydrox-

yphenylhydantoin (HPH) could be enhanced 200 fold by the substitution of Phe

159 with Ala. In this context, a working model for the enantioselectivity of HYD

is proposed here. As shown in Fig. 2, the HYD substrate binding pocket resembles

a barrel with a lid and consists of the recognition sites for the amide group (the

barrel bottom) and the functional R group of the substrate (the lid). The functional

release of the product from the enzyme occurs only if the chiral portion of the

substrate is complementarily coordinated with the active amino acid residues in the

hydrophobic core.

A key feature for enzymes of industrial application is their thermostability.

Thanks to the information acquired on molecular structures, comparison of a

thermostable HYD and HYD from a mesophilic microbe Burkholderia pickettii

Figure 2. Schematic illustration of a proposed model for the enantioselectivity of hydantoinase. (1) The

substrate binding pocket of HYD resembles a barrel with the lid open. (2) The model substrate is guided

into the barrel with the amide group downwards. (3) The amide group of the model substrate fits into

the barrel bottom (the amide recognition site), while the functional R group is correctly positioned and

oriented to make close contact with the lid. As a result, the lid closes and the conformational change

leads to the establishment of intimate interactions between the substrate and the protein’s active site

602 YUN-PENG CHAO ET AL.

(HYDbp) has been made possible (Xu et al., 2003), providing an insight into

the molecular basis of HYD thermostability. Despite sharing moderate sequence

homology, the two HYDs have high similarity as regards their overall and catalytic

site structures. The amino acid composition of the substrate recognition site is

however less conserved, which accounts for the difference in substrate specificity.

Moreover, it indicates that many factors including aromatic residues in the interior of

the structure, salt bridges and hydrogen-bonding interactions, and oxygen-sensitive

residues on the protein surface likely contribute to the varying thermostability

of HYD.

3. PURIFICATION AND IMMOBILIZATION

Progress in the practical application of proteins in industry is usually limited by

their production levels and purification processes. Overproduction of microbial

HYDs in E. coli is prone to the formation of insoluble inclusion bodies. To solve

this problem, a systematic study was undertaken to investigate the possible factors

affecting the solubility of HYD from Agrobacterium radiobacter NRRL B11291

(HYDar) (Chao et al., 2000). It was found that an active form of HYDar was

produced at low temperature and by coproduction of DnaJ/DnaK. In addition, by

fusion with thioredoxin HYDar’s solubility could be further improved with the

escort of DnaJ/DnaK. Meanwhile, high levels of production of a soluble fusion

protein comprising HYDbs and maltose binding protein (MBP) was reported (Kim

et al., 2000). However, a similar approach constructing a C-terminal fusion of

MBP to HYDar yielded the contrary: most of the MBP-fused protein aggregated

(unpublished data).

The fusion of affinity tags to a target protein usually makes the work of protein

purification easier. The N-terminal fusion of L-HYD of Arthrobacter aurescens to

MBP permitted a one-step purification of the hybrid protein using an amylase resin

(Pietzsch et al., 2000). However, the purification of authentic (unmodified) proteins

generally requires a tedious procedure consisting of multiple chromatographic steps.

To simplify this, we have recently developed a method using a single chromato-

graphic step for the purification of the recombinant HYDar (Huang et al., 2003).

After the recovery of cell-fee extract, HYDar without any affinity tags was purified

to high yield and high purity via a series of steps comprising heat treatment, alcohol

precipitation and chelating Sephacel chromatography.

The technique of confining enzymes into a defined space is a very useful approach

in many bioprocesses that require the use of enzymes since it facilitates enzyme

reutilization without the need for purification. Indeed, enzyme immobilization offers

advantages including the easy separation of the latter from the reaction products,

the elevation of enzyme concentrations per unit volume, and even the enhancement

of enzyme stability and activity. Recently, we have explored the utilization of the

chitin binding domain (ChBD) as an affinity tag to retain HYDar on chitin (Chern

and Chao, 2005). ChBD of chitinase A1 from Bacillus circulans WL-12 comprises

45 amino acids and exhibits remarkably high specificity for chitin (Hashimoto et al.,

HYDANTOINASES 603

2000). Compared to its unbound counterpart, immobilized HYDar exhibited higher

tolerance to heat and reached a half-life of 270 hr at 45



C. Notably, HYDar immobi-

lized on chitin could be reused 15 times to achieve conversion yields exceeding

90% using 100-mM HPH as the substrate.

Nevertheless, most of the HYDar produced in E. coli remains aggregated though

its solubility can be enhanced upon escort by molecular chaperons and the use of low

temperatures as mentioned above. Thus, in order to achieve the immobilization of

poorly soluble enzymes it is necessary to renature the insoluble proteins, a frequently

laborious, inefficient and costly process. A novel method based on artificial oil

bodies (AOBs) has recently been proposed (Chiang et al., 2006) to achieve protein

refolding and immobilization in one step. By fusing HYDar to the C terminus of

oleosin, a plant oil body storage protein (Frandsen et al., 2001), a hybrid protein

was obtained in the form of inclusion bodies in E. coli representing 30% of the total

cellular protein content. Without the need for addition of a denaturant, the insoluble

fusion protein combined with plant oil was readily incorporated into AOBs upon

sonication. It was found that in addition to its higher thermal resistance, HYDar

immobilized on AOBs could be maintained stable for at least 15 days when stored at



C. When applied repeatedly in a bioconversion process, the immobilized HYDar

gave a conversion yield exceeding 80% over 7 cycles. One added advantage of this

method is that it allows easy recovery of immobilized enzymes from the top of

a solution simply by performing a brief centrifugation. The essence of this novel

method for enzyme immobilization is outlined in Fig. 3.

It is well recognized that HYDar mediates the hydrolysis of HPH to N-carbamoyl-

D-hydroxyphenyl glycine (CpHPG), an intermediate which can be converted to

D-p-hydroxyphenylglcine (D-HPG) (Olivieri et al., 1979). D-HPG is of great

commercial value for its potential as a precursor of semi-synthetic antibiotics. Thus,

the successful achievement of HYD immobilization presents an important and

fundamental step in this bioconversion process.

Figure 3. Schematic illustration of the immobilization of hydantoinase on oil bodies. (1) The recovery

of the oleosin-fused hydantoinase in the form of inclusion bodies is mixed with plant oil and subjected

to sonication. (2) Hydantoinases immobilized on the surface of the oil bodies are removed from the top

of the aqueous phase after a brief centrifugation. (3) Oil bodies are fed into the buffer solution to initiate

the bioconversion reaction

604 YUN-PENG CHAO ET AL.

4. PRODUCTION OF D- OR L-AMINO ACIDS

In principle, enantiomerically pure D- or L-amino acids can be readily obtained

from racemic D,L-5-monosubstituted hydantoins in a two-step reaction directed

by the combined activity of stereoselective HYD and N-carbamoylase (Fig. 4).

This process for the production of optically pure amino acids has received consid-

erable interest by industry. Of particular importance, the production of D-HPG,

a precursor used in the synthesis of cephalosporins and obtained from D,L-HPH,

was first reported using A. radiobacter NRRL B11291 carrying D-HYD and

D-carbamoylase (Olivieri et al., 1981). To increase the efficiency of this process, a

recombinant E. coli strain overexpressing these two genes was constructed and used

for D-HPG production (Chao et al., 1999). A 5-fold increase in D-HPG productivity

was achieved by this approach compared to production by A. radiobacter NRRL

B11291. Moreover, the flux control coefficient of D-carbamoylase for this two-step

reaction was determined to be 0.8, suggesting that this enzymatic reaction is the

rate-determining step (Chao et al., 1999). This analysis suggests the possibility of

optimising D-HPG production by modifying the ratio of the two enzyme activities

in the reaction. Accordingly, the two respective genes from A. radiobacter NRRL

B11291 were cloned and expressed separately under the control of the tac promoter

(Chao et al., 2000). With a mixed population consisting of various proportions of

recombinant E. coli strains producing each individual enzyme, an optimal produc-

tivity of D-HPG was obtained with the ratio of D-carbamoylase to D-HYD activity

ranging between 1 and 2.

The production of L-amino acids using recombinant E. coli strain has also

been developed (Wilms et al., 2001). Similarly, the balanced activity of L-HYD,

L-carbamoylase, and racemase was investigated and established. Application to

the reaction process using the engineered E. coli cells gave a 6-fold increase in

productivity compared to that by the whole cell from A. aurescens.

Figure 4. Production of optically pure amino acids by the combined action of hydantoinase and

N-carbamoylase. (1) Hydrolysis of D,L-5-monosubstituted hydantoin by hydantoinase leads to the

production of the corresponding carbamoyl amino acids (carbamoyl-AA). (2) With the liberation of CO

deamination of carbamoyl-AA by steroselective N-carbamoylase leads to the production of optically

pure amino acids

HYDANTOINASES 605

5. FUTURE PERSPECTIVES

The relatively untapped potential of unusual amino acids in the pharmaceutical

industry will provide the incentive for the development of HYD-based production

processes. Rapid advances in the development of this production process can be

expected given the continuing efforts in expanding the substrate spectrum of HYD,

altering its enantioselectivity, and the improvement of its activity and thermosta-

bility. All of these goals can be achieved by the approach of directed evolution

(May et al., 2000) and the rational design of protein structures based on the

comparative analysis of molecular structures (Cheron et al., 2004). Meanwhile, the

availability and efficiency of preparative methods for immobilized enzymes and

immobilized whole cells should greatly increase the competitiveness and feasibility

of this production process. The initial approach of the immobilization of recom-

binant E. coli with k-carrageenan for D-HPG production proved unsatisfactory

(Chao et al., 1999). Owing to the low solubility of the substrate, the occurrence

of substrate fouling on the gel surface made the immobilized cells inappropriate

for reuse. Recently, ChBD-based cell immobilization has been explored by the

construction of a tripartite gene fusion consisting of lpp-ompA-ChBD in E. coli

(Wang and Chao, 2005). The result showed that ChBD-displaying cells exhibited

highly specific and stable binding to chitin within a wide range of pH (5–8) and

temperatures 15–37



C. It will be very interesting to investigate the potential of

this approach by fixing the whole cell on a chitin surface for D-HPG production.

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SUBJECT INDEX

 jelly-roll, 43, 72

(/

structure, 24

(/

structure, 43

(/

TIM-barrel fold, 3, 4, 8, 11, 13, 70, 601

(1 → 2), 148

(1 → 4), 143, 148

(1 → 6), 147

(2 → 1), 150

(2 → 6), 150, 151

-1, 3-glucans, 46

-amylase, 3, 10, 22, 24, 26, 31

-amylase, 3–10, 12–14, 21–23, 26–28, 31, 33,

38, 148, 149, 450, 453, 521

-chaconine, 131

-configuration, 22

-configuration, 8, 22

-D-fructofuranose, 150

-D-galacturonic acid, 99

-D-glucosidase, 36, 131, 133, 143

-fructofuranosidase, 150, 151

-galactosidase, 142, 143, 450

-glucosidase, 42, 43, 52, 130, 143

-glucosidase, 5, 8, 11, 12, 28, 143, 144, 148

-helices, 41, 105, 108, 109, 185, 221, 224, 364,

571, 572

-limit dextrin, 22

-limit dextrin, 24

-L-rhamnosidase, 117

-L-rhamnosidase, 117, 118, 120, 122, 123,

131, 132

-nitrophenyl--L-rhamnopyranoside

(pNPR), 124

-propeller, 43, 151, 254, 255, 505, 507, 511,

512, 515

(R)-(-)-glycidyl butyrate, 295

(R-(-)-mandelic acid, 534, 542

(R S-2-bromo hexanoic acids, 295

(S-ibuprofen, 542

(S-phenyllactic acid, 542

-sheet, 72, 106, 109, 199, 253, 270, 272, 382,

571, 572

-terpinyl esters, 285

-xylosidase, 66, 67, 74, 84

(/

-barrel, 4, 10, 11, 72

(/

-barrel, 13

(/

-barrel, 3, 4, 7–12, 20, 71

(/

-barrel catalytic domain, 11, 148

1, 10-phenanthroline, 229, 449

1, 5-dimethyl 2-piperidone, 542

16S rRNA, 422

1-butanol, 233, 326, 328, 329

-fructofuranosyl-nystose, 150–152

1-kestose, 150–152

1-propanol, 92, 326, 328, 329

2, 6-dimethoxyphenol, 462

2’-5’ branch linkage, 405, 411

23S rRNA, 422

2-O-lauroyl lactic acid, 294

2-O-palmitoyl lactic acid, 294

2-O-stearoyl lactic acid, 294

2-phenylethanol, 133

3’ to 5’ exonuclease, 382–386, 388, 389, 392

3’ hydroxyl group, 427

3-phosphoglyceraldehyde, 100

4, 5 unsaturated oligogalacturonate, 102, 103

4-hydroxybenzoic acid, 471

4-methylumbelliferyl--L-rhamnoside, 125

4-O-methylglucuronic, 73, 75, 84

4-t-butylcyclohexyl acetate, 285

5’ to 3’ exonuclease, 383, 384, 388

5’ phosphate group, 423, 427

5’-deoxyribose phosphatase activity, 383

5-keto-4-deoxy-uronate, 100

6-APA, 583, 584, 586–588, 591, 592

6-kestose, 152

/ hydrolase fold, 85, 90, 253, 254, 268, 270,

272, 343

// sandwich fold, 85

607