FEMS Microbiology Letters 35 (1986) 99-105 99 Published by Elsevier

FEM 02451

Enzymes metabolizing dimethylamine, trimethylamine and trimethylamine N-oxide in the yeast

Sporopachydermia cereana grown on amines as sole nitrogen source

(Flavin nucleotides; carbon monoxide; amino mono-oxygenases; trimethylamine N-oxide reductase)

Dav id Whi t f ie ld and Peter J. Large *

Department of Biochemistry, University of Hull. Hull. HU6 7RX. U.K.

Received 7 February 1986 Accepted 3 March 1986

1. S U M M A R Y

Sporopachydermia cereana, an ascosporogenous yeast, grew on dimethylamine, trimethylamine or trimethylamine N-oxide as sole nitrogen sources and produced mono-oxygenases for dimethyl- amine and trimethylamine that were significantly more stable than the corresponding enzymes found in Candida utilis. N o trimethylamine mono- oxygenase activity was found in S. cereana grown on dimethylamine. In cells grown on trimethyl- amine N-oxide (but not on the other nitrogen sources), evidence for an enzyme metabolizing the N-oxide, possibly an aldolase, but more probably a reductase was obtained. All these activities showed a similar requirement for the presence of FAD or FMN in the extract buffer during isola- tion to retain activity. Amine mono-oxygenase activities showed a similar sensitivity to inhibitors, including proadifen hydrochloride and carbon monoxide as the corresponding enzymes in C. utilis. The tr imethylamine N-oxide-dependent oxidation of N A D H was more sensitive to inhibi-

* To whom correspondence should be addressed.

tion by EDTA, N-ethylmaleimide and fl-phenyl- ethylamine than the mono-oxygenases, and less sensitive to KCN, and activity was significantly higher with N A D P H than was observed with the 2 mono-oxygenases.

2. I N T R O D U C T I O N

4370 of the 461 yeast species tested by van Dijken and Bos [1] were able to grow on mono-, di- and tri-methylamine as sole nitrogen source. Although trimethylamine N-oxide was not tested, yeasts are known which will also use this com- pound as sole nitrogen source [2,3]. The oxidation systems for di- and trimethylamine have been investigated in Candida boidinii [3,4] and C. utilis [5], and shown to be N A D H - or NADPH-depen- dent mono-oxygenases (Eqns. 1 and 2)

(CH3)3N + 02 + N A D ( P ) H + H +

--' (CH3)2NH + NAD(P) ÷ + H C H O + H 2 0

(CH3)2NH + 02 + N A D ( P ) H + H ÷

--~ CH3NH 2 + NAD(P) ÷ + H C H O + H 2 0 (2)

0378-1097/86/$03.50 ~) 1986 Federation of European Microbiological Societies

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These enzymes were both present in C. utilis, irrespective of whether dimethylamine or tri- methylamine was the nitrogen source. They were very unstable and inhibited by carbon monoxide, suggesting that they were microsomal haemopro- teins [3-5]. Although C. utilis [5], C. boidinii [3] and Candida strain WY-3 [2] can also grow on trimethylamine N-oxide as sole nitrogen source, no evidence could be found for an enzyme (re- ductase or demethylase) that could metabolize this compound. A search was undertaken to find another, more stable, source of these activities to enable them to be characterized further. Initial studies indicated that the ascosporogenous yeast S. cereana [6] was a potential alternative. In this organism, for the first time in yeasts, evidence has been obtained for the reduction of trimethylamine N-oxide (Eqn. 3)

(CH 3 )3NO + N A D ( P ) H + H +

(CH3)3N + NAD(P)+ + H 2 0 (3)

3. MATERIALS A N D M ETHODS

3.1. Materials Chemicals, enzymes and inhibitors were ob-

tained from the sources described previously [5].

3. 2. Growth and maintenance of the cultures S. cereana CBS6644 and C. utifis NCYC321

were grown and maintained as described for C. boidinii by Haywood and Large [7], the carbon source being 55 mM glucose and the nitrogen source 30 mM di- or trimethylamine or trimethyl- amine N-oxide. Growth was measured as ab- sorbance at 663 nm. Cells were harvested by centrifugation at 10000 x g for 10 min at 4°C, and washed with 50 mM potassium phosphate buffer pH 7.0. Harvesting took place when A063 was 1.5-2.0 ( S. cereana ) or 1.0-1.5 ( C. utilis ).

3.3. Preparation of cell-free extracts Cells were suspended in an equal volume of

ice-cold 50 mM potassium phosphate buffer pH 7.0 containing 20 p,M FAD (buffer A), unless otherwise stated, and passed once through a French pressure cell operated at 7.85 MPa. Un-

broken cells and cell debris were removed by centrifugation at 40000 x g for 10 min at 4°C.

3. 4. Preparation of washed microsomal suspensions The crude extract was centrifuged for 60 min at

105000 x g at 4°C to produce a microsomal pel- let. The pellet was resuspended in twice the origi- nal volume of ice-cold buffer A and recentrifuged as before to produce a washed microsomal pellet. This was resuspended in buffer A to give a protein concentration of 1-2 mg/ml . The preparation was used immediately.

3.5. Assay of enzyme actwities The spectrophotometric assay method (a) of

Green and Large [5] was employed with a sub- strate concentration of 2 mM for dimethylamine- HCI and 4 mM for trimethylamine-HCl or tri- methylamine N-oxide-HCl (unless otherwise stated).

3. 6. Determination of kinetic parameters" K~, °v and Vma x values were obtained by varying

the amine substrates and their concentration and plotting reciprocal rates against reciprocal con- centrations. Lines were fitted by linear regression [8]. Variations, especially in Vm, x values, were observed between some preparations for some amines, but values were within one order of mag- nitude. Representative data like those in Table 2 are to allow comparison between substrates.

3. 7. Chemical determinations Protein was determined by the method of Brad-

ford [91.

3.8. Inhibition experiments Inhibitors were preincubated with enzyme in

the standard assay mixture for 5 min in the ab- sence of substrate and cofactor before the reaction was started by addition of substrate.

4. RESULTS

4. I. General properties of dimethylamine and tri- methylamine mono-oxygenases in S. cereana

S. cereana grew on amines as sole nitrogen

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source rather more slowly than did C. utilis, and needed also to reach a higher culture density for maximal enzyme activity. For maximum activity of the enzymes, S. cereana was harvested after 24 h growth on dimethylamine or trimethylamine, compared with 16 h for C. utilis. When trimethyl- amine N-oxide was the nitrogen source, S. cereana

cultures took 48 h to reach maximum activity levels. The endogenous rate of N A D H oxidation was also 2- to 4-fold higher in S. cereana than in C. utilis. Moreover yields of microsomal protein per g dry weight of cells were lower in S. cereana

(29 m g / g dry wt.) compared with C. utilis (56 m g / g dry wt.). One important difference from the Candida species was that no trimethylamine mono-oxygenase activity could be detected in mi- crosomes from S. cereana grown on dimethyl- amine.

The general properties of dimethylamine and trimethylamine mono-oxygenases in S. cereana

showed a few differences from the corresponding enzymes in C. utilis. The pH optima for the oxida-

tion of dimethylamine and trimethylamine by washed microsomes were respectively 7.5 and 8.0, compared with 7.0 and 7.5 for C. utilis and 6.7 and 7.0 for C. boidinii [3]. While dimethylamine mono-oxygenase specific activity was similar or slightly lower in S. cereana than in C. utilis,

trimethylamine mono-oxygenase specific activity in the former was usually about 2-fold higher than in C. utilis.

Substitution of 0.13 mM NADPH for 0.13 mM N A D H in the standard assay produced rates that were only 8-9% of those observed with N A D H as cofactor, so the latter coenzyme was used in all further experiments.

4.2. Effect o f addition o f f lavin nucleotides on mono-oxygenase activity

As previously observed in C. utilis [5], micro- somes of S. cereana prepared in the absence of FAD showed a much reduced capacity to oxidize dimethylamine and trimethylamine and this also proved true for reduction of trimethylamine N-

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8O O O

a

< 60 U. ~r =L 0 04 v

40 >,

,~ 2o 2,

0 I I I I 5 10 15 20

C o n c n of FAD in buf fer " d u r i n g p r e p a r a t i o n (jaM)

Fig. 1. Effect on recoveries of activities of dimethylamine and trimethylamine mono-oxygenases of the presence of various concentrations of FAD during the preparation of washed microsomal suspensions from S. cereana. O, Dimethylamine mono-oxygenase activity; o, trimethylamine mono-oxygenase activity.

too

8o

io, 1o 20 30 40 50 6o

T i m e ot 4 0 " C (min)

>,6O

4O "6

g o E 2O

I1.

Fig. 2. Effect of heating at 40°C on the enzyme activities of washed microsomal preparations from S. cereana. After vari- ous lengths of time at 40°C, samples were rapidly cooled in ice and the residual activity measured in the standard assays: o, dimethylamine mono-oxygenase; A trimethylamine mono- oxygenase; m, trimethylamine N-oxide reductase. 100% activity was, respectively, for the three enzymes: 160, 90, and 49/.tmol NADH oxidized/min/g microsomal protein.

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oxide . T h e p re sence o f 20 # M F A D in the bu f f e r

t h r o u g h o u t the m i c r o s o m a l i so la t ion p r o c e d u r e re-

su l ted in a 5- fo ld inc rease in the speci f ic ac t iv i t ies

o f di- and t r i - m e t h y l a m i n e m o n o - o x y g e n a s e s (Fig.

1). In the a b s e n c e o f F A D in the i so la t ion buf fe r ,

no t r i m e t h y l a m i n e N - o x i d e - d e p e n d e n t o x i d a t i o n Nitrogen Temper-

o f N A D ( P ) H cou ld be de tec ted . F M N was j u s t as source for ature Dimethyl- e f fec t ive as F A D in these expe r imen t s . N o synerg- growth (°C) amine

istic ef fec t was o b s e r v e d w h e n F M N was a d d e d to mono-

the assay cuve t t e s of ex t rac t s p r e p a r e d in the oxygenase

p r e s e n c e o f 5 - 1 0 t tM F A D . Trimethyl- 35 43.5

A d d i t i o n o f 5 - 2 0 t tM F A D to the assay mix- amine 40 57.0 N-oxide 45 36.0

ture on ly s t i m u l a t e d the e n z y m i c ra te w h e n the

m i c r o s o m e s had been p r e p a r e d in the p r e sence of Trimethyl- 35 49.5 amine 40 57.0 low ( 0 - 5 t tM) c o n c e n t r a t i o n s o f F A D . If con -

45 41.0 c e n t r a t i o n s o f 1 0 / ~ M F A D or h igher were p re sen t

d u r i n g the i so la t ion p rocedu re , a d d i t i o n of F A D Dimethyl- 35 54.0 tO the assay m i x t u r e caused a c o n c e n t r a t i o n - d e - amine 40 60.0

45 50.0 p e n d e n t i nh ib i t i on o f ac t iv i ty o f up to 20%. T h e

Table 1

Half-lives of dimethylamine mono-oxygenase, trimethylamine mono-oxygenase and trimethylamine N-oxide reductase activi- ties at various temperatures in microsomes prepared from cells grown on all three nitrogen sources

Half-life (rain) of enzyme activity

Trimethyl- Trimethyl- amine amine mono- N-oxide oxygenase reductase

30.0 9.0 8.0

31.5 18.0 4.5

40.5 14.0 10.0

Table 2

Kinetic parameters for secondary and tertiary amine mono-oxygenases of S. cereana

Values are repre~ntative data from 2-4 separate preparations. Secondary amines were assayed at pH 7.5, tertiary amines at pH 8.0. n.d., No activity detected.

Amine substrate Nitrogen source for growth:

Dimethylamine Trimethylamine Trimethylamine N-oxide

K m Vm. ~ K~ V,~,a ~ K,. Vm,,x (~M) (U/g (#M) (U/g (#M) (U/g

protein) protein) protein)

Secondary amines Dimethylamine 14 186 17 95 18 357 Diethylamine 27 143 28 143 25 215 Dipropylamine 73 101 180 123 100 95 Dibutylamine 36 85 25 54 n.d. 0 Di-isobutylamine 255 32 274 40 178 36 Methylethylamine 14 150 18 243 15 449 Methylpropylamine 183 88 79 94 76 209 Methylisopropylamine 335 145 257 138 80 121 Methylbutylamine 15 68 16 217 17 38 Methyloctylamine 53 49 53 83 50 98 Methylethanolamine 335 239 106 230 269 291 Ethylpropylamine 54 79 50 168 55 300

Tertiary amines Trimethylamine n.d. 0 15 50 14 60 Dimethylethylamine 2 65 16 136 8 112 Diethylmethylamine 5 54 6 154 7 205 Triethylamine n.d. 0 6 47 3 59

activity reconstituted by addition of FAD to the assay mixture was always significantly lower than in microsomes prepared by breaking open cells in the presence of FAD.

4.3. Effect o f heat on mono-oxygenase and tri- rnethylamine N-oxide reductase activities

The effect of exposing microsomes for various lengths of time to temperatures of 35°C, 40°C and 45°C was investigated, using cells grown on di- or trimethylamine or trimethylamine N-oxide as nitrogen source. Representative data for 40°C for microsomes of cells grown on trimethylamine N- oxide are shown in Fig. 2, and typical half-lives for the activities from the various preparations are summarized in Table 1. The reason for the ap- parent decreased stability of dimethylamine mono-oxygenase at 35°C compared with 40°C is not known. At all temperatures trimethylamine mono-oxygenase activity was less stable than tri- methylamine N-oxide reductase, while dimethyl-

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amine mono-oxygenase was the most stable of all. The activities from S. cereana are thus signifi- cantly more stable than the corresponding activi- ties in C. utilis (half-lives at 35°C: 0.4 min for trimethylamine monooxygenase activity and 1.7 min for dimethylamine mono-oxygenase, [5]). Sim- ilarly, microsomal preparations from S. cereana stored in ice for 24 h retained 70-75% of both mono-oxygenase activities, whereas C. utilis pre- parations lost 80-100% of both activities.

4.4. Kinetic data Apparent K,,, and Vmax values for amine sub-

strates of the S. cereana mono-oxygenases are summarized in Table 2. The K~ pp value for NADH could not be determined, due to the high rate of substrate-independent NADH oxidation by mi- crosomal preparations.

4. 5. Inhibitor studies The pattern of inhibition of dimethylamine and

Table 3

Effect of various inhibitors on dimethylamine mono-oxygenase, trimethylamine mono-oxygenase and trimethylamine N-oxide reductase activities

For experimental details see section 3.8.

Inhibitor Concentration Percentage inhibition of activity

(mM) Dimethyl- Trimethyl- Trimethyl- amine amine amine mono- mono- N-oxide oxygenase oxygenase reductase

trans-2- Phenyl- cyclopropylamine 0.25 99 97.5 95.1

Pargyline hydrochloride I. 16 19.3 0 41.7 Aminoacetonitrile 0.10 0 29.4 8.1 fl-Phenylethylamine 0.05 0 8.2 45.9 Benzylamine 0.05 0 17.4 45.1 Hydroxyammonium chloride 0.5 16.3 16.1 0 1,10-Phenanthroline 5.0 38.7 79.4 73.9 2,2'-Bipyridyl 0.5 0 0 0 Disodium EDTA 2.5 0 0 17.4 Potassium cyanide 0.5 59.8 83.7 37.3 n-Octylamine 1.0 40.0 51.0 76.5 Ethanol 10.0 0 65.2 47.5 Proadifen-HC1 (SKF 525-A) 0.5 96.4 68.2 100.0 Metyrapone 1.0 12.0 55.8 0 N-Ethylmaleimide 1.25 7.4 1.9 53.0 2-Mercaptoethanol 0.5 80.0 0 100.0 Dithiothreitol 0.05 18.6 0 20.0

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trimethylamine mono-oxygenase and trimethyl- amine N-oxide reductase activities in microsomal preparations by a wide range of possible com- pounds is shown in Table 3. For the mono- oxygenases, the results are similar to those ob- served previously with C. utilis [5], with the excep- tion of the effect of carbon monoxide. Examina- tion of the kinetics of carbon monoxide inhibition showed that it was uncompetitive for both mono- oxygenases, and the K i values were 3.8 #M for dimethylamine mono-oxygenase and 2.2 #M for trimethylamine mono-oxygenase, compared with figures for the C. utilis system [51 of 7.5 #M and 0.35 #M, respectively. Thus trimethylamine mono-oxygenase activity of S. cereana is signifi- cantly less sensitive to carbon monoxide.

4. 6. Tr imethy lamine N-ox ide reductase activity

Microsomal preparations from S. cereana

grown on trimethylamine N-oxide as sole nitrogen source catalysed the oxidation of NADH or NADPH on addition of trimethylamine N-oxide. NADPH was 33% as effective as NADH in reduc- ing trimethylamine N-oxide when tested at 0.13 mM in the standard assay. The K m value for trimethylamine N-oxide was 17.6/~M and the lima x value was 58.4 U / g protein. The reductase activity was not detectable in extracts of cells grown on dimethylamine or trimethylamine. The pH opti- mum for trimethylamine N-oxide reduction was 8.0. The inhibition data of Table 3 show dif- ferences of inhibition from the trimethylamine and dimethylamine mono-oxygenases for fl-phe- nylethylamine, benzylamine, hydroxyammonium chloride, EDTA, KCN, metyrapone, mercapto- ethanol and N-ethylmaleimide. Inhibition by carbon monoxide was non-competitive with a K~ of 1.0 #M. Despite the differences, the data sup- port the hypothesis that the reductase is also a haemoprotein-dependent activity.

5. DISCUSSION

The evidence from this work with S. cereana clearly indicates that secondary and tertiary amines are oxidized by different mono-oxygenases, since trimethylamine mono-oxygenase activity was ab-

sent from dimethylamine-grown cells. The fact that both mono-oxygenases were formed in C utilis and C. boidinii during growth on dimethyl- amine, together with the inability to separate the two activities meant that it was impossible to prove for certain that two separate enzymes were involved, although their markedly different sensi- tivities to heat did indicate this [5].

The two mono-oxygenases of S. cereana are not only more stable than those of C. utilis, but the trimethylamine mono-oxygenase is also signifi- cantly less sensitive to inhibition by carbon mono- xide. Otherwise, the differences between the two systems are relatively slight. S. cereana micro- somes may thus be a better starting point for purifying these activities.

The demonstration of a trimethylamine N- oxide-dependent oxidation of NAD(P)H in S. cereana tends to confirm an earlier suggestion [5,10] that trimethylamine N-oxide is catabolized by reduction to trimethylamine. The stimulation by flavins is in agreement with observations on other N-oxide reductase systems in mammalian liver microsomes [11,12]. This is the first report of the occurrence of an enzyme catalysing the metabolism of trimethylamine N-oxide in a yeast. The identity of the product of the reaction re- mains to be confirmed. The trimethylamine N- oxide-dependent oxidation of NADH could in theory be due to the concerted action of a non- oxidative trimethylamine N-oxide aldolase (Eqn. 4) followed by dimethylamine mono-oxygenase

(CH3)3NO --~ (CH3)2NH + HCHO (4)

(Eqn. 2), producing a combined stoichiometry equivalent to Eqn. 3. Trimethylamine N-oxide al- dolase activity occurs in bacteria [13-15], and has also been reported in fish muscle microsomes [16]. Three pieces of evidence make the suggestion that an aldolase is involved less likely in our view. First, if an aldolase converted the N-oxide directly to dimethylamine, there would be no role for trimethylamine mono-oxygenase in S. cereana

grown on the N-oxide. Secondly the higher activ- ity of trimethylamine N-oxide reduction with NADPH compared with dimethylamine mono- oxygenase would be difficult to explain, and thirdly oxygen consumption during oxidation of tri-

methylamine N-oxide by washed mlcrosomes pro- ceeds at the same rate as dur ing oxidat ion of t r imethylamine (results not shown) and not slower as it would if a preceding step were rate-limiting, nor faster as it might if the N-oxide were giving rise to d imethylamine, which is more rapidly oxidized than t r imethylamine (Table 2 and Fig. 2). However, conf i rmat ion of the nature of the reac- t ion involving t r imethylamine N-oxide is obvi- ously necessary, by purif icat ion of the enzyme(s)

involved.

A C K N O W L E D G E M E N T S

We thank Geoff Haywood for drawing the diagrams. This work was supported by the Science and Engineering Research Counci l (Gran t No. G R / C / 5 4 7 7 7 ) whose suppor t is gra teful ly

acknowledged.

R E F E R E N C E S

[1] van Dijken, J.P. and Bos, P. (1981) Arch. Microbiol. 128, 320-324.

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[2] Yamada, H., Kishimoto, N. and Kumagai, H. (1976) J. Ferment. Technol. 54, 726-737.

[3] Green, J. and Large, P.J. (1983) Biochem. Biophys. Res. Commun. 113, 900-907.

[41 Green, J. and Large, P.J. (1984) J. Gen. Microbiol. 130, 1947 - 1959.

[5] Green, J. and Large, P.J. (1984) J. Gen. Microbiol. 130, 2577-2588.

[6] Rodrigues de Miranda, L. (1978) Ant. van Leeuwenhoek 44, 439-483.

[7] Haywood, G.W. and Large, P.J. (1981) Biochem. J. 199, 187-201.

[8] Brook, D.F. and Large, P.J. (1976) Biochem. J. 157, 197- 205.

[9] Bradford, M.M. (1976) Anal. Biochem. 72, 248-254. [101 Large, P.J. and Green, J. (1984) in Microbial Growth on

C l Compounds, Proceedings of Fourth International Symposium (Crawford, R.L. and Hanson, R.S., Eds.) pp. 155-164. American Society for Microbiology, Washing- ton, DC.

111] Dajani, R.M., Gorrod, J.W. and Beckett, A.H. (1975) Biochem. Pharmacol. 24, 109-117.

[121 Sugiura, M., lwasaki, K. and Kato, R. (1976) Mol. Pharmacol. 12, 322-334.

[13] Large, P.J. (1981) in Microbial Growth on C 1 Com- pounds, Proceedings of Third International Symposium (Dalton, H., Ed.) pp. 55-69. Heyden and Son, London.

[14] Myers, P.A. and Zatman, L.J. (1971) Biochem. J. 121, 10P. [15] Large, P.J. (1971) FEBS Lett. 18, 297-300. [161 Parkin, K.L. and Hultin, H.O. (1982) FEBS Lett. 139,

61-64.