**1. Introduction**

Gel electrophoresis of enzymes is a very useful and powerful analytical method, which is at present widely used in many distinct fields of both biological and medical sciences and successfully applied in many different fields of human activity. The tremendous expansion of this methodology is mainly due to its simplicity and its high ability to separate both isoenzymes and alloenzymes, which have proven to be very useful genetic markers. The most important step of enzyme electrophoresis is the detection of native enzymes on electrophoretic gels; it means the procedure of obtaining electropherograms, or zymograms. Detection of enzymes on electrophoresis gels means the visualization of gel areas occupied by specific enzyme molecules after their electrophoretic separation. From this point of view, sometimes the testing of an enzyme-coding DNA sequence for expression of catalytically active enzyme is performed by zymograms, where the use of this technique for this purpose is very effective, cheap, and time saving. The number of applications of zymogram techniques for testing cloning enzyme-coding genes for their expression at the protein level is growing (Pfeiffeer-Guglielmi et al., 2000; Lim et al., 2001; Okwumabua et al., 2001). An absolute prerequisite for this is the specific and sensitive zymogram technique suitable for detection of the enzyme inside the gel and the use of the appropriate substrates.

The zymograms has been used to detect a variety of oxidoreductases (Bergmeyer, 1983), including isoenzymes (Jeng & Wayman, 1987) and to classify various genera of yeast based upon the relative mobility of the activity bands produced by selected enzymes (Goto & Takami, 1986; Yamasaki & Komagata, 1983). Electrophorezed gels are placed in a staining solution containing a reduced substrate such as an alcohol, oxidized cofactor such as NAD+ or NADP+, a dye such as nitroblue tetrazolium, and an electron acceptor-donor such as phenazine methosulphate. At the location of the appropriate enzyme catalyzing oxidation of substrate and reduction of cofactor, a dark-purple band appears as a result of the precipitate that forms upon reduction of the dye.

Polyacrylamide Gel Electrophoresis an Important Tool for the

glucose that was incubated for 12 h at same other conditions.

different fractions were kept at -70 °C for further studies.

**2.3 Gel electrophoresis** 

**2.4 Enzymatic assays** 

**2.2 Preparation of purified fractions** 

Detection and Analysis of Enzymatic Activities by Electrophoretic Zymograms 249

yeast-peptone-glucose-agar (YPGA) (Bartnicki-García & Nickerson, 1962) was used for strain maintenance, spore collection and mycelium growth. Aerobic mycelium growth was also carried out on salt minimal medium (Alvarado et al. 2002) added with 0.1% (w/v) peptone (sMMP). As a carbon source we added D-glucose (1% w/v) or glycerol (1.0% v/v) or ethanol (2.0% v/v) or *n*-decanol (1.0% v/v) or *n-*pentane (1.0% v/v) or *n-*decane, (1.0% v/v) or *n-*hexadecane (1.0% w/v) or naphthalene (0.5% w/v) or anthracene (0.5% v/v) or phenanthrene (0.5% w/v) or pyrene (0.5% w/v). Liquid cultures (600 ml) inoculated with spores at a final cell density of 5 x 105/ml were propagated in 2-l Erlenmeyer flasks and incubated in a reciprocating water bath at 28°C for 22 h at 125 rpm for all substrates except

Mycelium of 22 h of incubation was harvested by filtration and exhaustively washed with cold sterile-distilled water; mycelial mass was suspended in 15 ml of 20 mM Tris-HCl pH 8.5 buffer containing 1 mM phenylmethanesulphonyl fluoride (previously dissolved in ethanol). Approximately 20 ml of cells was mixed with an equal volume of glass beads (0.45- 0.50 mm diameter) and disrupted in a Braun model MSK cell homogenizer (Braun, Melsungen, Germany) for four periods of 30 sec each under a CO2 stream. The homogenate (crude extract) was centrifuged at 4,300*g* for 10 min in a J2-21 Beckman rotor in a Beckman JA-20 centrifuge to remove cell walls and unbroken cells, a 1 ml sample of the supernatant was saved. The rest of the supernatant (low speed supernatant) was centrifuged at 31,000*g* for 20 min in a 70Ti Beckman rotor in a Beckman L8-80 ultracentrifuge and samples of 1 ml of the supernatant was saved; the resulting pellet (mitochondrion rich sample) was resuspended in 2 ml of buffer and saved. The rest of the supernatant was high-speed centrifuged at 164,500*g* for 45 min in a 70Ti Beckman rotor at 4ºC in a Beckman L8-80 ultracentrifuge; the supernatant (cytosolic fraction) was put aside, and the pellet, the mixed membrane fraction (MMF), was resuspended in 2 ml and saved. In all cases samples of

The slab gels were 1.5 mm-thick contained 6% (w/v) acrylamide/4% (w/v) bisacrilamide, loaded with the cytosolic fraction of each culture and run in the mini-gel system manufactured by Bio-Rad. The continuous buffer system described by Laemmli (1970) without SDS (native conditions) was used to run for 2.5 h at 80 V. The *Rm* values were calculated as the ratio of the distance migrated by the stained band divided by the distance migrated by tracking dye; standard deviation was calculated with Excel from three independent experiments and each experiment was made by triplicate on each substrate.

All enzyme assays were carried out in a final volume of 1 ml and incubated for different times at 25 ºC. NAD+-dependent ADH activity was assayed in the oxidative direction according to Bergmeyer (1983). The enzymatic assays contained 25 mM Tris-HCl (pH 8.5), 2 mM NAD+ or NADP+, cell-free extract (100-200 µg protein), and 100 mM of the substrate (1R, 2S)-*cis*-1,2-di-hydro-1,2-naphthalene-diol. The reaction was started by dihydrodiol addition, and reduction of NAD+ or NADP+ was monitored by the increase in absorbance at

Previously we have investigated the use of zymogram staining of native electrophoretic gels as an initial approach to the identification of carbonyl reductase activities against both aliphatic (Silva et al., 2009; Zazueta et al., 2008) and aromatic hydrocarbons (Durón et al., 2005; Zazueta et al., 2003) in *Mucor circinelloides* YR1, an indigenous fungus isolated from petroleum contaminated soil.

Oil spills sometimes occur during routine operations associated with the exploration and production of crude oil. Crude oils vary widening in composition depending on factors such as source bed type and generation temperatures (Hunt, 1979). Biodegradation rates for crude oils will vary due to differences in composition, as reflected by hydrocarbon class distribution: saturates, aromatics, and polars, and the amount *n-*alkanes *versus* branched and cyclic alkanes within the saturated hydrocarbon class (Cook et al., 1974). In nature exist many types of microorganisms useful in the biodegradation processes of contaminant compounds (Atlas, 1995), such as the polycyclic aromatic hydrocarbons (PAH's) that are persistent soil contaminants and many of which have toxic and carcinogenic properties (Hyötyläinen and Oikari, 1999; Cerniglia, 1997).

In bacterial aerobic degradation of aromatic compounds, reactions of metabolic pathways generally lead to the formation of aromatic intermediates containing two hydroxyl constituents, which are subsequently ring-cleaved by excision dioxygenases (Neidle et al., 1992). In many catabolic pathways the formation of such intermediates is carried out by two successive enzymatic steps namely dihydroxylation of the polyaromatic substrate to produce *cis*-diols followed by dehydrogenation (Harayama & Timmis, 1989). The ring hydroxylation is catalyzed by multi-component dioxygenases, while the dehydrogenation is catalyzed by *cis*diol-dehydrogenases. In mammalian tissues the enzyme dihydro-diol dehydrogenase (DD, EC 1.3.1.20) exists in multiple forms (Hara et al., 1990; Higaki et al., 2002) and catalyses the NADP+-linked oxidation of *trans*-dihydro-diols of aromatic hydrocarbons to the corresponding catechols (Penning et al., 1999). Studies on the metabolism of aromatic hydrocarbons by fungi are limited, nevertheless have been shown to posses the ability to metabolize aromatic compounds (Auret et al., 1971; Ferris et al., 1976) and the aryl oxidative enzymes of fungi appear to be similar to monooxygenases of hepatic microsomes (Cerniglia & Gibson, 1977; Ferris et al., 1976). Smith & Rosazza (1974) have also presented evidence that naphthalene is metabolized to 1-naphtol by six different genera of fungi.

In this work we analyze the cytosolic fraction of YR-1 strain by electrophoretic zymograms, methodology that there is not described in the literature for the NADP+-dependent dihydrodiol dehydrogenase (DD) activities. We analyze all the activity bands corresponding to proteins with DD activity present in an enzymatic extract in only one lane of the electrophoretic gel. Our results show eleven different DD activity bands, five of them are constitutive, DD1-5, since they appears when the strain is growth on glucose, and the others six are induced by different compound added to the culture media as a sole carbon source. Some biochemical-enzyme characteristics as pH, optimal temperature, cofactor dependence, substrate specificity and the effect of cations, EDTA and pyrazole were investigated for DD activities when YR-1 strain was grown in naphthalene as sole carbon source.
