dehydrogenase activity in yeast experiment

19. Investigation of dehydrogenase activity in yeast

• 00:22 Which molecules act as a hydrogen acceptors during aerobic respiration?
• 00:29 How will you adapt this method to investigate dehydrogenase activity in yeast?

redox indicator: methylene blue (0.05g/100cm 3 )

yeast suspension (100g/dm -3 )

30 o C Water bath

Cork for test tube

10 cm 3 syringe

1 cm 3 syringe

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Article Contents

1 introduction, 2 isoenzymes of yadh, 3 substrate specificity, 4 steady-state kinetic mechanism, 5 pre-steady-state kinetics, 6 primary structure, 7 the active site, 8 mutations in the yeast enzyme, 9 binding of coenzymes, 10 chemical mechanism, 11 hydride transfer, acknowledgements.

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The three zinc-containing alcohol dehydrogenases from baker's yeast, Saccharomyces cerevisiae

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Vladimir Leskovac, Svetlana Trivić, Draginja Peričin, The three zinc-containing alcohol dehydrogenases from baker's yeast, Saccharomyces cerevisiae , FEMS Yeast Research , Volume 2, Issue 4, December 2002, Pages 481–494, https://doi.org/10.1111/j.1567-1364.2002.tb00116.x

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This review is a summary of our current knowledge of the structure, function and mechanism of action of the three zinc-containing alcohol dehydrogenases, YADH-1, YADH-2 and YADH-3, in baker's yeast, Saccharomyces cerevisiae . The opening section deals with the substrate specificity of the enzymes, covering the steady-state kinetic data for its most known substrates. In the following sections, the kinetic mechanism for this enzyme is reported, along with the values of all rate constants in the mechanism. The complete primary structures of the three isoenzymes of YADH are given, and the model of the 3D structure of the active site is presented. All known artificial mutations in the primary structure of the YADH are covered in full and described in detail. Further, the chemical mechanism of action for YADH is presented along with the complement of steady-state and ligand-binding data supporting this mechanism. Finally, the bio-organic chemistry of the hydride-transfer reactions catalyzed by the enzyme is covered: this chemistry explains the narrow substrate specificity and the enantioselectivity of the yeast enzyme.

Yeast alcohol dehydrogenase (EC 1.1.1.1) is a member of a large family of zinc-containing alcohol dehydrogenases. The primary structures of 47 members of this family have been determined and aligned, and an evolutionary tree has been constructed, assuming a divergent evolution from a common ancestral gene [ 1 ]. In this way, it was possible to identify four divergent groups of alcohol dehydrogenases in this family: vertebrates, plants, eukaryotic microorganisms and prokaryotic bacteria. Baker's yeast ( Saccharomyces cerevisiae ), a member of the third group, has three isoenzymes of alcohol dehydrogenase: YADH-1, YADH-2, and YADH-3. YADH-1 is the constitutive form that is expressed during anaerobic fermentation [ 2 ]. YADH-2 is another cytoplasmic form, which is repressed by glucose [ 3 ], and YADH-3 is found in the mitochondria [ 4 ]. YADH-1 accounts for the major part of alcohol dehydrogenase activity in growing baker's yeast.

The structure, function and mechanism of action of yeast alcohol dehydrogenase have been reviewed three decades ago [ 5 , 6 ]. The purpose of this article is to update the subject and to review novel data on the structure, function and mechanism of action of the isoenzyme YADH-1; this isoenzyme will be abbreviated as YADH throughout the text. The steady-state kinetic constants are presented in the nomenclature of Cleland [ 7 ].

Yeast alcohol dehydrogenase was one of the first enzymes to be purified and isolated [ 8 ]. If the steady-state kinetic properties of the ADH isoenzymes are compared, a large degree of similarity is detected. Table 1 shows the steady-state kinetic constants for the three isoenzymes of YADH, isolated from baker's yeast.

Steady-state kinetic constants of yeast ADH isoenzymes with ethanol and acetaldehyde as substrates, at pH 7.3, 30°C a

a Calculated from the data of Ganzhorn et al. [ 9 ].

It is evident that YADH-1 and YADH-3 have very similar kinetic characteristics, while YADH-2 has a much higher substrate specificity for ethanol ( V 1 / K B ) and acetaldehyde ( V 2 / K P ), and much lower Michaelis constants with ethanol ( K B ) and acetaldehyde ( K P ). Recently, the kinetic characterization of YADH-1 and YADH-2 has been extended by measuring their specificity constants ( V 1 / K B ) for a number of long-chain alcohols and diols. It was found that for all alcohols, normalized rates with YADH-2 were about three-fold faster than with YADH-1 [ 10 ].

At neutral pH, the equilibrium is shifted far to the left ( Table 2 ).

Steady-state kinetic constants for the oxidation of various alcohols at neutral pH

a Calculated from the data of Dickinson and Monger [ 11 ], at pH 7.0, 25°C.

b Calculated from the data of Schöpp and Aurich [ 12 ], at pH 8.0, 25°C.

c Calculated from the data of Trivić and Leskovac [ 13 ], at pH 7.0, 25°C.

d Calculated from the data of Trivić and Leskovac [ 14 ], at pH 7.0, 25°C.

e Calculated from the data of Chen and Huang [ 15 ], at pH 8.2, 25°C.

f K eq = V 1 K iQ K P /( V 2 K iA K B ).

Substrate specificity of YADH is restricted to primary unbranched aliphatic alcohols, and any branching in the side chain diminishes the activity of the enzyme and lowers its efficiency. In addition, the enzyme also shows activity towards secondary alcohols. Table 2 presents the steady-state kinetic constants for various alcoholic substrates and Table 3 shows the steady-state constants for various carbonyl substrates of the yeast enzyme.

Steady-state kinetic constants for the reduction of various carbonyl substrates at neutral pH

b Calculated from the data of Trivić and Leskovac [ 13 ], at pH 7.0, 25°C.

c Calculated from the data of Leskovac et al. [ 16 ], at pH 9.0, 25°C.

d Calculated from the data of Trivić et al. [ 17 ], at pH 8.9, 25°C.

e Calculated from the data of Leskovac et al. [ 16 ], at pH 7.0, 25°C.

Ethanol is by far the best substrate of the yeast enzyme. Methanol is a very poor substrate of YADH; the methanol activity of the enzyme at pH 8.8 is only 0.07% of its ethanol activity under identical conditions. The enzyme is able to oxidize methanol by NAD + to formaldehyde and NADH, but the enzymatic reaction is very complex due to interference of numerous side reactions [ 18 ].

Allyl and cinnamyl alcohol are, however, excellent substrates; kinetic constants for the latter alcohol are: V 1 =133 s −1 and V 1 / K B =29 mM −1 s −1 , at pH 8.2, 25°C [ 19 ]. ( S )-(+)-Butan-2-ol is a much better substrate than ( R )-(−)-butan-2-ol ( V 1 =1.0 and 0.05 s −1 , and V 1 / K B =18 and 0.8 M −1 s −1 , respectively, at pH 7.3, 30°C) [ 20 ]. 4-Methyl-1-pentanol ( V 1 =7 s −1 , pH 8.2) is a much better substrate than 2-methyl-1-propanol ( V 1 =0.2 s −1 , pH 7.3) or 3-methyl-1-butanol ( V 1 =0.3 s −1 , pH 8.2) [ 19 , 20 ].

It was reported that glycerol, glyceraldehyde and acetol are poor substrates of YADH [ 21 ], whereas benzyl alcohol and benzaldehyde are extremely poor substrates of this enzyme [ 20 , 22 ]. It has also been reported that p -chlorobenzyl alcohol and p -methoxybenzyl alcohol are slowly oxidized by NAD + in the presence of YADH [ 23 ]. 2-Chloroethanol, 2-fluoroethanol, 2,2,2-trifluoroethanol, propargyl alcohol, glycidol and polyethylene glycol are no substrates of the yeast enzyme [ 24 ].

Chloroacetaldehyde is an excellent substrate of YADH ( Table 3 ), while 2-chloroethanol is not oxidized by NAD + , which makes the reaction 2 essentially irreversible [ 16 ]. p -Nitroso- N , N -dimethylaniline (NDMA) is readily reduced by NADH, in the presence of YADH ( reaction 4 ); the primary product of this reaction, the corresponding hydroxylamine, is transformed into a quinonediimine compound by the loss of a molecule of water. The last compound is reduced non-enzymatically by NADH to p -amino- N , N -dimethylaniline [ 17 , 25 ]. YADH has a weak aldehyde dehydrogenase activity; it is able to catalyze an irreversible oxidation of acetaldehyde to acetic acid with NAD + , with an apparent k cat =2.3 s −1 and V / K =34 M −1 s −1 , at pH 8.8, 22°C [ 26 ].

Free acetaldehyde is a true substrate for alcohol dehydrogenase [ 27 ], and gem -diol is probably a true substrate for aldehyde dehydrogenase activity of YADH [ 26 ].

Yeast alcohol dehydrogenase catalyzes the chemical reactions described by Eq. 1 . Numerous investigations of the steady-state kinetic mechanism of the yeast enzyme have been conducted by several authors [ 9 , 11 , 28–36 ]; they have led to the conclusion that the yeast enzyme follows the steady-state random mechanism on the alcohol side, and a steady-state ordered mechanism on the aldehyde side of the catalytic cycle, with primary aliphatic alcohols and aldehydes ( Scheme 1 ).

The Brändén mechanism.

Eq. 6 satisfies the results obtained for the reduction of acetaldehyde and butyraldehyde in predicting a linear reciprocal equation, in which the K iQ , V 2 / K Q and V 2 K iQ / K Q constants are independent of the nature of the aldehyde ( Table 3 ).

The kinetic mechanism in Scheme 1 is compatible with deuterium isotope effects on maximal rates reported for ethanol, D V 1 =1.8, D V 1 / K A =1.8, and D V 1 / K B =3.2 [ 39 ], propan-1-ol, D V 1 =3.7 [ 40 ], butan-1-ol, D V 1 =3.7 [ 41 ], and propan-2-ol, D V 1 =2.2 around neutrality [ 13 ]. With ethanol, the effect on D V 1 / K A was smaller than on D V 1 / K B , suggesting that NAD + binds before ethanol; the still significant size of D V 1 / K A is probably due to dissociation of NAD + from the ternary complex [ 39 ].

With propan-2-ol and acetone, the kinetic mechanism is steady-state random in both directions [ 13 ]. A similar kinetic mechanism probably holds for most branched and secondary alcohols [ 34 ].

Pre-steady-state kinetic studies provide the numerical values of the rate constants in the mechanism. The pre-steady-state kinetics of yeast alcohol dehydrogenase has been studied with the help of the KINSIM and FITSIM computer programs of Frieden [ 42–44 ].

These computer software packages can simulate the reaction progress curves and calculate the individual rate constants therefrom ( Table 4 ). The magnitudes of the individual rate constants in Scheme 1 were calculated from reaction progress curves in both directions, keeping the concentration of reactants at such a level that dissociation of NAD + from the central complex was prevented, and therefore excluding the rate constants k 11 – k 14 ( Table 4 ) [ 45 ].

Thermodynamics of the yeast alcohol dehydrogenase reaction, at pH 7.0, 25°C [ 45 ]

a Data in parentheses are from the steady-state kinetic measurements of Dickinson and Dickenson [ 31 ], at pH 7.0, 25°C.

b Taken from Northrop [ 46 ].

c Calculated from the equilibrium constant: k 4 / k 3 =( k 41 / k 31 )/( k 42 / k 32 ).

d Calculated from the Haldane relationship: K eq = V 1 K iQ K p /( V 2 K iA K B ).

One can see from Table 4 that the magnitudes of rate constants obtained from the computer simulation of reaction progress curves [ 45 ] and from the steady-state kinetics [ 31 ] are very similar, the differences reflecting only the different enzyme preparations.

In the horse liver enzyme, a large conformational change of the enzyme is triggered when the coenzyme binds, well documented both in structural terms [ 47 ] and by kinetic methods [ 48 ]. Recently, Northrop has reported that moderate pressure increases the capture of benzyl alcohol ( V 1 / K B ) in YADH-catalyzed oxidation of this alcohol with NAD + , by activating the hydride transfer step [ 49 ]. This means that the collision complex for hydride transfer (*E·NAD + ) has a smaller volume than the free alcohol plus the capturing form of the enzyme (E·NAD + ) [ 46 ]. This was a direct experimental proof for the isomerization step in the yeast enzyme, which enabled the estimation of the equilibrium constant k 41 / k 31 [ 75 ]; using this value, it was possible to calculate the equilibrium constant k 42 / k 32 ( Table 4 ).

Inspection of data in Table 4 clearly shows that, in the forward direction (oxidation of ethanol at neutral pH), the rate-limiting step is not the chemical reaction ( k 9 ), but the dissociation of NADH from the EQ-complex ( k 7 ). Likewise, NAD + dissociates much faster from the EA-complex ( k 2 ) than NADH dissociates from the EQ-complex ( k 7 ).

YADH-1 is a tetramer, composed of four identical subunits; each subunit consists of a single polypeptide chain with 347 amino acids, with a molecular mass of 36 kDa [ 47 ]. Each subunit has one coenzyme-binding site and one firmly bound zinc atom, which is essential for catalysis [ 50 , 51 ]; the catalytic domain provides the ligands to this zinc atom: Cys-46, His-67 and Cys-174. The second zinc atom/subunit is liganded in a tetrahedral arrangement by four sulfur atoms from the cysteine residues 97, 100, 103 and 111; this zinc atom only has a structural role [ 52 ].

Table 5 shows the primary structures of the three isoenzymes of YADH [ 4 , 53–55 ]. The alignment of amino acid residues for all 47 members of the ADH family was made progressively rather than pairwise [ 1 ].

Primary structure of the three isoenzymes of yeast alcohol dehydrogenase

The amino acid sequences of horse liver alcohol dehydrogenase and YADH-1 are homologous, and the homology amounts to 25% of the amino acid residues [ 5 ]. YADH-1 has been crystallized, but only preliminary crystallographic studies have been reported [ 56 ]. The three-dimensional structure of horse liver alcohol dehydrogenase in several binary and ternary complexes with coenzymes, substrates and inhibitors has been solved at high resolution [ 47 ]. The tertiary structures of liver and yeast enzyme are highly similar and able to accommodate extensive sequence changes between the enzymes [ 57 ].

Analogous to the liver enzyme, the subunits of the yeast enzyme are probably divided into two domains: the catalytic domain and the coenzyme-binding domain. The two domains are unequal in size; the catalytic domain contains 3/5 of all amino acids, whereas the coenzyme-binding domain contains the remaining 2/5 of the amino acids. The domains are separated by a cleft, containing a deep pocket which accommodates the substrate and the nicotinamide moiety of the coenzyme. One domain binds the coenzyme and the other provides ligands to the catalytic zinc, as well as to most of the groups that control substrate specificity [ 47 ].

Since the liver and yeast enzymes are homologous, molecular modeling of the yeast enzyme can approximate the structure of one subunit, but not yet the quaternary arrangement [ 57 ].

Fig. 1 shows a model of the active site of the yeast enzyme, drawn schematically after a model obtained in a molecular graphics display system by Plapp et al. [ 58 ]. The 3D-model of the active site of YADH provides an illustration of the main working machinery of the yeast enzyme. In order to perform catalysis, the active site of the enzyme has to bind a molecule of substrate and a molecule of coenzyme in a productive mode, and, subsequently catalyze a hydride-transfer reaction between them.

Model of the active site of yeast alcohol dehydrogenase, drawn schematically after Plapp et al. [ 58 ].

The adenosine-binding site is easily accessible from solution, whereas the nicotinamide-binding site is situated at the center of the molecule, buried deep inside the protein [ 47 ]. Numerous amino acid residues in the primary structure of the enzyme are involved in substrate and coenzyme binding and in catalysis ( Table 6 ).

Positions of residues that participate in enzymatic functions of the yeast enzyme (adopted from Jörnval et al. [ 57 ])

7.1 Substrate-binding pocket

The inner wall of the pocket is lined with hydrophobic side chains from the residues Trp-57, Trp-93, Asn-110, Leu-132, Tyr-140, Thr-141, Met-294, Ala-296, and Ile-318, which are from the same subunit as the zinc ligands. The substrate-binding site near zinc is narrow, because access is limited by Trp-93 and Thr-48.

The voluminous amino acid side chains of Trp-57, Trp-93 and Met-294 make the substrate-binding pocket in the yeast enzyme much more narrow than the corresponding pocket in the horse liver enzyme.

7.2 Ligands to the active-site zinc

At the bottom of the substrate-binding pocket, a zinc atom is coordinated to three protein ligands: two thiolates from Cys-46 and Cys-174, and one imidazole nitrogen of His-67. The other imidazole nitrogen of His-67 is hydrogen-bonded to a carboxylic group of Asp-49. A carboxylic group of Glu-68 is also in close proximity to the active-site zinc atom. Asp-49 and Glu-68 are the residues conserved in all known zinc-dependent alcohol dehydrogenases; instead of being inner-shpere ligands to the zinc, both amino acids are situated in the second sphere. The only polar groups in the pocket close to zinc are the zinc ligands, the nicotinamide moiety of the coenzyme, and the side chain of Thr-48.

7.3 Nicotinamide ring

The nicotinamide ring binds in a cleft in the interior of the protein, close to the center of the molecule. On one side the ring interacts with Thr-178, Leu-203 and Met-294. The other side faces the active site, and is close to the catalytic zinc atom and the sulfur ligands of Cys-46 and Cys-174.

The oxygen atom of the carboxamide group is hydrogen-bonded to the main-chain nitrogen atom of Val-319. The nitrogen atom of the carboxamide group is hydrogen-bonded to the carboxyl oxygens of Val-292 and Ser-317. The side chain of Thr-178 helps to keep the nicotinamide ring of the nucleotide in the correct stereochemical position for hydride transfer ( Fig. 5 ); Thr-178 is conserved in all known homologous alcohol dehydrogenases.

Stereospecificity of YADH catalysis. NADH binds anti , presenting Re -hydrogen (H Re ) to acetaldehyde lying above the coenzyme in this diagram. For clarity, Thr-178 is not shown; the methyl group of this side chain lies below and to the left of the nicotinamide behind Leu-203 (reproduced from Weinhold et al. [ 64 ], with permission of the corresponding author).

7.4 The proton-relay system

The chemical mechanism of action of alcohol dehydrogenase [ 36 ].

7.5 Binding of the coenzyme

The coenzyme is bound to the apoenzyme by numerous secondary valence forces. Important amino acid residues are: Asp-223, which is hydrogen-bounded to AMP-ribose, His-47, forming a salt bridge with AMP-orthophosphate, and Leu-203, forming a hydrogen bond to NMN-orthophosphate.

The three yeast ADH genes have been cloned and described [ 4 , 54 , 55 ]. Therefore, it was possible to change individual amino acids in the primary structure of YADH-1 via site-directed mutagenesis and isolate a quantity of mutated enzymes ( Table 7 ).

Steady-state kinetic constants for YADH mutants, with ethanol and acetaldehyde as substrates, determined at pH 7.3, 30°C

ND=not determined.

a Determined at pH 8.2, 25°C.

In recent years, a number of genetically engineered mutants of YADH-1 were isolated and kinetically characterized, principally by Plapp and his co-workers. Most of these mutations involve amino acids which are intimately involved in the binding of substrates and in catalysis, and provide information about the general principles concerning the function of the catalytic residues. Table 7 shows the steady-state kinetic properties of all YADH mutants described so far participating in substrate binding and in catalysis.

8.1 Substrate-binding pocket (Met-294, Trp-57, Trp-93)

An exchange of Leu for Met-294, on the edge of the substrate-binding pocket, has very little influence on the steady-state kinetic properties of the enzyme with ethanol or acetaldehyde. On the other hand, the Met294Leu mutant has a 10-fold lower catalytic activity ( V 1 ) with butan-1-ol, indicating that the C4-atom of butan-1-ol is in a close contact with Met-294, whereas the shorter ethanol is not [ 9 ]. An exchange of Met or Leu for Trp-57 decreases the catalytic efficiency ( V 1 / K B ) with ethanol only three- to four-fold, whereas an exchange of Ala for Trp-93 decreases the catalytic efficiency 300-fold; with an enlargement of the substrate-binding pocket in the latter case, the enzyme acquires weak activity with branched-chain alcohols (2-methyl-1-butanol, 3-methyl-1-butanol) and benzyl alcohol [ 19 , 20 ].

8.2 Ligands to the active-site zinc (Asp-49, Glu-68)

The carboxylate group of Asp-49 is hydrogen-bonded to His-67, which in turn coordinates the active-site zinc; in addition, the carboxylate group of Glu-68 is in the vicinity of the active-site zinc. If Asn is substituted for Asp-49 or Gln for Glu-68, a negative charge is removed from the vicinity of the active-site zinc; these substitutions reduce the catalytic efficiency with ethanol ( V 1 / K B ) 1000 times and 100 times, respectively, and the catalytic constant ( V 1 ) 40 times. These reductions in activity were interpreted by an increased electrostatic potential near the active-site zinc, due to removal of negative charges; as a consequence the activity is decreased by hindering isomerizations of enzyme–substrate complexes [ 39 ].

8.3 The proton-relay system (Thr-48, His-51)

An exchange of Ser for Thr-48 does not interrupt the hydrogen bonding in the proton relay system and, as expected, the activity of the Thr48Ser mutant is very similar to that of the wild-type. The double mutant Thr48Ser:Trp93Ala and the triple mutant Thr48Ser:Trp57Met:Trp93Ala show decreased activities that are obviously due to removal of bulky tryptophan residues from the substrate-binding pocket [ 20 ]. An exchange of Cys or Ala for Thr-48 disrupts the hydrogen bonding in the relay system and, as expected, renders the enzyme inactive [ 58 ].

The role of His-51 in catalysis has been tested by replacing it with glutamine or glutamic acid [ 58 , 61 ]. These residues have an appropriate size to form the hydrogen bond with the 2′-hydroxyl group of the nicotinamide ribose; thus, binding of the coenzyme in the mutant enzymes could resemble binding in the wild-type enzyme. On the other hand, a glutamine residue would not be able to participate in base catalysis, whereas a glutamate residue could accept a proton. Plapp et al. [ 58 ] have found that a wild-type enzyme has a distinct p K a value of 7.7 in the pH-profile for the V 1 / K B function. Replacement of His-51 with Gln or Glu reduces the value of V 1 / K B 13-fold and 60-fold at pH 7.3, respectively; in addition, the p K a value of 7.7 in the pH profile of the V 1 / K B function is abolished in both cases. These results were interpreted by a mechanism in which the amino acid residue in the mutant enzyme hinders the deprotonation of alcohol through the proton relay system [ 58 ]. On that interpretation, these results are consistent with the role of His-51 in the proton relay system, where it participates as a base.

Fig. 2 summarizes the steady-state and ligand-binding data relevant for the binding of coenzymes to the free enzyme.

pH profiles for the binding parameters of coenzymes to the free enzyme; rate constants k 7 and k 8 , as in Scheme 1 (adopted from Leskovac et al. [ 34 ]).

The dissociation constant of the E·NAD + complex for the yeast enzyme, K E,NAD + , is practically pH-independent; on the other hand, the dissociation constant of the E·NADH complex, K E,NADH , decreases with lower pH-value over three apparent p K a values (6.6, 8.0 and 9.0). The association rate constant for the binding of NADH to the free enzyme ( k 8 ) decreases in alkaline over a single p K a value 7.8, while the dissociation rate constant for the E·NADH-complex ( k 7 ) is almost pH-independent, from pH 6.5 to 9.0.

In recent years, a number of genetically engineered mutants of YADH-I, with mutations in the adenylate-binding pocket, have been isolated and kinetically characterized, principally by Plapp and his co-workers ( Table 8 ). The following general conclusions may be drawn from the kinetic data shown in Table 8 .

Steady-state kinetic constants for YADH mutants in the adenylate binding pocket, with ethanol and acetaldehyde as substrates, determined at pH 7.3, 30°C

b Alignment of amino acids according to Table 5 . Ala200 is an insertion in the yeast enzyme with respect to other members of the alcohol dehydrogenase family of enzymes; therefore, this residue is not counted in the primary structure that follows after this residue [ 1 ].

9.1 Adenine site substitutions (Ser-198, Gly-224, Gly-225)

Gly224Ile and Gly225Arg mutants have only modest effects on coenzyme binding and other kinetic constants, but the Ser198Phe mutant significantly decreases its affinity for coenzymes and turnover numbers.

9.2 Adenosine–ribose binding (Asp-223)

The Asp223Gly:Gly225Arg double mutant shows a decrease in all kinetic parameters, but uses NAD(H) and NADP(H) with about the same efficiency.

9.3 Pyrophosphate binding (His-47, Ala-200, Leu-203, Gly-204)

Mutation of the residues Ala-200, Leu-203 or Gly-204 decreases all kinetic parameters significantly, suggesting that these amino acids are essential for the binding of the pyrophosphate moiety of the coenzyme. On the other hand, substitution of His-47 by the basic amino acid Arg decreases the catalytic activity with NAD(H) only modestly.

9.4 Nicotinamide–ribose binding (Ser-269)

The Ser269Ile mutant decreases its turnover numbers by 350-fold.

Studies of the mutants in the adenylate-binding site of the enzyme show that several amino acid residues at the proposed adenylate-binding site of the enzyme are important for coenzyme binding and formation of productive ternary complexes. The Asp223Gly:Gly225Arg double mutant was the only mutant that uses NAD(H) and NADP(H) with about the same efficiency; this result suggests that conversion of the coenzyme specifically requires multiple substitutions [ 62 ]. Mutations of amino acids Leu-203 and Thr-178 have been performed in order to locate the structural determinants of the high stereospecificity of the enzyme for the coenzyme NAD(H) [ 64 ].

Primary structure, tertiary structure and point mutations in the yeast enzyme, outlined in the preceding sections, strongly suggest that the integrity of the proton relay system is indispensable for the activity of the enzyme.

Based on this integrity of the relay system, which is maintained throughout the catalytic cycle, Cook and Cleland [ 60 ] have proposed the chemical mechanism of action for the yeast enzyme as shown in Scheme 2 .

In this mechanism, B and P represent alcohol and ketone, and k 3 , k 4 , k 5 and k 6 represent hydride-transfer steps; X is an intermediate with the stoichiometry of an alkoxide, and k 1 and k 2 are the steps in which a proton is transferred from B to a group on the enzyme to give X , and similarly for the reverse process.

An assignment of appropriate p K a values to all dissociation forms of the enzyme in Scheme 2 was founded on studies of the pH dependence of the steady-state kinetics and ligand-binding parameters [ 14 , 16 , 26 , 33–36 , 65 ], as outlined below.

Table 9 shows the macroscopic p K a values calculated from the pH profiles of the maximal rates ( V 1 ) and the specificity constants ( V / K ) with various substrates.

Macroscopic p K a values and pH-independent limiting constants in various YADH-catalyzed reactions (adopted from Leskovac et al. [ 36 ])

Table 10 presents the p K a values calculated from the pH profiles of binding constants ( K i ) for competitive dead-end inhibitors.

Macroscopic p K a values and pH-independent constants for ternary complexes of YADH with competitive dead-end inhibitors a

a Calculated from the data of Leskovac et al. [ 35 ].

The specificity constants V / K with ‘nonsticky’ substrates, such as propan-1-ol, propan-2-ol, NDMA, DACA and acetone, provide information on catalytically active groups in enzyme–coenzyme complexes [ 66 ], if the pH profiles of V / K are fitted to initial-rate equations appropriate to the mechanism in Scheme 2 [ 36 ]. In this way, the p K 1 (8.0) and p K 5 (7.9–8.0) values in Scheme 2 were estimated. From the binding of azide, a dead-end inhibitor competitive with alcohols, the value for p K 1 (7.9) was confirmed; from the binding of acetamide, a dead-end inhibitor competitive with aldehydes, the values for p K 4 (8.3) and p K 5 (7.9) were estimated.

pH profiles for the V 1 function provide information on catalytically active groups in the productive ternary enzyme·NAD + ·alcohol-complex [ 66 ]. In this way the p K 2 value was estimated (8.3), from the pH profiles of V 1 with butan-1-ol and propan-2-ol.

An indirect estimation provided the value of p K 3 (8.3) [ 36 ].

The chemical mechanism of action, presented in Scheme 1 , can be drawn entirely in terms of the proton relay system, as is shown in Fig. 3 ; in Fig. 3 , however, the Thr-48 residue was omitted from the relay for the sake of simplicity. The key feature of Fig. 3 is that His-51 lies at the surface of the protein and thus can be deprotonated as in the conversion of HEAX to EAX or HEQP to EPQ, while reactants are bound and the state of protonation of molecules in the substrate-binding site is locked. Thus, HEAX can be deprotonated to EAX without preventing subsequent hydride transfer.

A different view on the chemical mechanism of action of yeast alcohol dehydrogenase has been presented by Brändén et al. [ 5 ]. These authors proposed that the Zn 2+ -bound water dissociates when the coenzyme NAD + is added; the remaining (OH) − deprotonates the alcohol, which is then bound to the Zn 2+ ion as the fourth ligand. Fig. 4 shows this dissociation in the proton relay system.

Recently, Nadolny and Zundel [ 67 ] have claimed experimental evidence supporting the above mechanism. These authors obtained Fourier-transform infrared (FTIR) spectra of various complexes of yeast alcohol dehydrogenase with NAD + and coenzyme analogs; from the FTIR spectra they concluded that, upon binding of NAD + to the enzyme, N1 of the coenzyme adenosine becomes protonated and the molecule of water in the active site dissociates to a hydroxyl anion. It was postulated that the positive charge is conducted from the zinc-bound water to histidine-51 and then further to the N1-atom of the adenine rest via the proton relay system through the protein. Thus the binding of NAD + to the enzyme shifts the equilibrium 1→2 in Fig. 4 to the right. The substrate, alcohol, is then deprotonated by the (OH) − bound to the Zn 2+ ion and forms the structure 3.

The experiments of Nadolny and Zundel [ 67 ] with the yeast enzyme were conducted at pH 7.5 and they do not explain the pH dependence of the enzyme activity. Further, the proposed mechanism lacks the explanation for the conductance of the positive charge from His-51 to adenine across a distance of approximately 7 Å.

One of the classical aspects of coenzyme binding to yeast alcohol dehydrogenase is the A-stereospecificity of the coenzyme [ 68 ]. YADH-catalyzed reactions are highly stereospecific; the enzyme catalyzes the transfer of the Re -hydrogen (pro- R or A-type) at the 4-position of NADH to the carbonyl carbon of the substrate ( Fig. 5 ).

The stereochemical fidelity of the hydride transfer reaction is very high, and YADH makes but one stereochemical ‘mistake’ every 7 000 000 turnovers. If the bulky side chain of Leu-203 is exchanged with Ala, the Leu203Ala mutant ( Table 7 ) makes one stereochemical ‘mistake’ every 850 000 turnovers with NADH, and every 450 turnovers with thio-NADH, which has a weaker hydrogen-bonding capacity. From this, it was concluded that the decrease in stereochemical fidelity comes from an increase in the transfer rate of the 4- Si -hydrogen of NADH. The nicotinamide ring of the coenzyme is kept in a correct position for hydride transfer mainly by hydrogen bonds between its amide group and Val-292 and Val-319, and the rotation of 180° around the glycosidic bond is obstructed mainly by the side chain of Leu-203 [ 64 ].

The main reaction catalyzed by alcohol dehydrogenase is, in principle, a very simple reaction. An alcohol group is oxidized by the removal of a proton from the hydroxyl group and by the transfer of a hydride ion from the adjacent carbon atom to NAD + . By analogy with the horse liver enzyme [ 47 ], we may assume that hydride transfer in the yeast enzyme occurs in a completely water-free environment. Direct transfer of a hydride ion is facilitated in a hydrophobic environment, where water is excluded. The positive charge on the nicotinamide ring is crucial for the enhanced binding of alcohol to the enzyme; insertion of the positive charge in this hydrophobic environment facilitates formation of the negatively charged alcoholate ion. The creation of an alcoholate ion greatly facilitates hydride transfer. The important role of the zinc atom in alcohol oxidation is to stabilize the alcoholate ion for the hydride-transfer step. In the reverse direction, zinc functions as an electron attractor, which gives rise to an increased electrophilic character of the aldehyde, consequently facilitating the transfer of a hydride ion to the aldehyde. Thus, the proposed mechanism is essentially electrophilic catalysis mediated by the active-site zinc atom.

The overall oxidation of alcohol to aldehyde involves the net release of one proton ( Eq. 1 ); the ultimate source of this proton is alcohol. The release of a proton from the bound alcohol occurs in the center of the enzyme molecule in a region that is inaccessible to solution; the proton is transferred by certain groups on the enzyme to the surrounding solution ( Fig. 1 ). Because water is not directly involved in the catalytic reaction, that is, no hydrolysis or hydration, there is no reason to suggest a role for a water molecule at the active site of YADH [ 36 ].

In catalysis, the molecules of the substrate and the nicotinamide ring of the coenzyme probably do not have fixed positions. The rearrangement of electron configuration on the carbon atom from the sp 2 hybridization in aldehyde to the sp 3 in alcohol, requires different pathways for hydride transfer and, consequently, different relative orientations [ 69 ].

Cha et al. [ 74 ] have demonstrated that, in this reaction, the exponents in Eq. 8 are 3.58 and 10.2 for the primary and secondary kinetic isotope effect, respectively, indicating significant breakdown of the semi-classical upper limit. For hydrogen tunneling to occur, the reactive carbon atoms have been brought close together so that the classical energy barrier is penetrated. Thus, it appears that hydrogen tunneling is an additional general phenomenon which facilitates the YADH catalysis [ 23 , 76–78 ].

Leskovac et al. [ 79 ] have studied the primary kinetic isotope effects and the internal thermodynamics of the YADH-catalyzed oxidation of 2-propanol- h 8 and 2-propanol- d 8 with NAD + ; the properties of this reaction were compared with non-enzymatic model redox reactions of N 1 -substituted-1,4( 1 H 2 )dihydronicotinamides and N 1 -substituted-1,4( 1 H 2 H)dihydronicotinamides with a number of various oxidizing agents. The kinetic and thermodynamic properties of the enzymatic reaction closely resemble the model hydride-transfer reactions which probably proceed via a linear transition state, and are very dissimilar from reactions which proceed via a bent transition state, suggesting that this particular enzymatic reaction has a linear transition state.

This work was financially supported by the Ministry of Science and Technology of the Republic of Serbia.

Abbreviations

yeast alcohol dehydrogenase, isoenzyme YADH-1

p -nitroso- N , N -dimethylaniline

sodium azide

N , N -dimethylamino- trans -cinnamaldehyde

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14 Dehydrogenase in yeast

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Investigating the Rate of Respiration ( AQA A Level Biology )

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Biology & Environmental Systems and Societies

Required Practical: Investigating the Effect on the Rate of Respiration

• A redox indicator is a substance that changes colour when it is reduced or oxidised
• They are used to investigate the effects of temperature and substrate concentration on the rate of anaerobic respiration in yeast
• These dyes can be added to a suspension of living yeast cells as they don’t damage cells
• Yeast can respire both aerobically and anaerobically, in this experiment it is their rate of anaerobic respiration that is being investigated
• Dehydrogenation happens regularly throughout the different stages of aerobic respiration
• The hydrogens that are removed from substrate molecules are transferred to the final stage of aerobic respiration, oxidative phosphorylation, via the hydrogen carriers NAD and FAD
• The enzyme dehydrogenase catalyses the production of reduced NAD in glycolysis
• When DCPIP or methylene blue are present they take up hydrogens from the organic compounds and get reduced instead of NAD
• Blue → colourless
• This means that the rate of colour change can correspond to the rate dehydrogenase would be working at and therefore, the rate of respiration in yeast
• The rate of respiration is inversely proportional to the time taken

Rate of respiration (sec -1 ) = 1 / time (sec)

• Glucose solution

Method - Temperature

• Add a set volume of yeast suspension to test tubes containing a certain concentration of glucose
• Put the test tube in a  temperature-controlled water bath and leave for 5 minutes to ensure the water temperature is correct and not continuing to increase or decrease
• Add a set volume of DCPIP to the test tube and start the stopwatch immediately
• This is subjective and therefore the same person should be assigned this task for all repeat experiments
• Repeat across a range of temperatures. For example, 30 o C, 35 o C, 40 o C, 45 o C
• For example, 0.1% glucose, 0.5% glucose, 1.0% glucose

Methylene blue or DCPIP is added to a solution of anaerobically respiring yeast cells in a glucose solution. The rate at which the solution turns from blue to colourless gives the rate of dehydrogenase activity.

Controlling other variables

• Volume of dye added : if there is more dye molecules present then the time taken for the colour change to occur will be longer
• Volume of yeast suspension : when more yeast cells are present the rate of respiration will be inflated
• Type of substrate : yeast cells will respire different substrates at different rates
• Concentration of substrate : if there is limited substrate in one tube then the respiration of those yeast cells will be limited
• Temperature : an increase or decrease in temperature can affect the rate of respiration due to energy demands and kinetic energy changes. The temperature of the dye being added also needs to be considered
• pH : a buffer solution can be used to control the pH level to ensure that no enzymes are denatured
• A graph should be plotted of temperature against time
• This means hydrogens are released by the reactions more quickly, hence the DCPIP accepts electrons/hydrogens more quickly until all molecules of DCPIP are reduced. This means that it will take less time to turn from blue to colourless

Limitations

• This experiment is not measuring the rate of dehydrogenase activity directly (through measuring the rate of substrate use or product made) but is instead predicting what the rate would be measuring the rate of electron / hydrogen release from the reactions of respiration
• Distinguishing the end of the reaction and the colour change is subjective and therefore one person should be used to attempt to control this

Although the DCPIP and methylene blue undergo a colour change from blue to colourless it is important to remember that the yeast suspension in the test tube may have a slight colour (usually yellow). That means when the dye changes to colourless there may still be an overall yellow colour in the test tube. If this is the case it can be useful to have a control tube containing the same yeast suspension but with no dye added, then you can tell when the dye has completely changed colour.

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Author: Alistair

Alistair graduated from Oxford University with a degree in Biological Sciences. He has taught GCSE/IGCSE Biology, as well as Biology and Environmental Systems & Societies for the International Baccalaureate Diploma Programme. While teaching in Oxford, Alistair completed his MA Education as Head of Department for Environmental Systems & Societies. Alistair has continued to pursue his interests in ecology and environmental science, recently gaining an MSc in Wildlife Biology & Conservation with Edinburgh Napier University.

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• Published: 08 May 1954

Evidence for the Occurrence of Glucose Dehydrogenase in Yeast

• A. R. FAHMY 1 &
• E. O'F. WALSH 1  

Nature volume  173 ,  page 872 ( 1954 ) Cite this article

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WHEN buffered suspensions of washed bakers' yeast are incubated anaerobically with methylene blue, dehydrogenase activity as measured by the rate of decolorization is greater with glucose than it is with other substrates. While investigating the action of nicotine on various dehydrogenase systems, including those of yeast, we found that, whereas nicotine inhibited dehydrogenase activity of Saccharomyces cerevisiae with glucose, activation resulted with glucose-6-phosphate as substrate.

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FAHMY, A., WALSH, E. Evidence for the Occurrence of Glucose Dehydrogenase in Yeast. Nature 173 , 872 (1954). https://doi.org/10.1038/173872a0

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Does temperature have an effect on the activity of dehydrogenase enzymes in yeast cells?

The Effect Of Temperature On Enzyme Activity

Hypothesis:   does temperature have an effect on the activity of dehydrogenase enzymes in yeast cells.

Aim:  The aim of this experiment is to show the relationship between temperature and the rate of activity of dehydrogenase enzymes in two different yeast types (‘Bakers’ and ‘Brewers’). We can then compare the results of the two different types of yeast to see which type is more affected by changes in temperature. Triphenyl tetrazolium chloride (TTC) is an artificial hydrogen acceptor, or redox indicator. When oxidised, TTC is colourless, but when it is reduced, TTC will form red, insoluble compounds called formazans. This colour change therefore shows the presence of active dehydrogenase enzymes in yeast cells. The temperature of the TTC solution and yeast suspension will affect the rate at which this colour change occurs, which in turn will show how the activity of dehydrogenase enzymes in different yeast cells changes.

Background Theory: Enzymes are tertiary protein structures made up of a single polypeptide chain. The polypeptide chain is folded into a precise shape, giving enzymes their specificity. Enzymes maintain this permanent shape by a range of bonds holding them together, including disulphide bridges, ionic bonds and hydrogen bonds. Enzymes are specific to the reactions that they catalyse, so the active site on an enzyme has a particular shape into which a specific substrate will fit. Enzymes can be described as globular proteins and are all biological catalysts. A catalyst is a substance which alters the rate of a chemical reaction without itself undergoing permanent change. As they are not altered they can be reused again and again. The active sites are responsible for the functioning of the enzyme. Anything which affects the three-dimensional shape of the protein molecule, will affect its ability to function.

Graph of activation energy.

Reactions involve breaking and remaking chemical bonds. For molecules to react, they first need sufficient energy, known as ‘activation energy’ needed to break the initial bonds. The higher the activation energy, the slower the reaction. Enzymes speed up reactions by lowering the activation energy required for a chemical reaction. This is brought about by the enzyme forming a complex with the substrate(s) for the reaction.

Substrate + Enzyme     →     Enzyme/substrate complex     →     Enzyme + Product

Once the products are formed, they are released and the enzyme is free to form a complex with another substrate. This process is summed up in the lock and key mechanism.

Lock and Key Mechanism - Within the structure of each enzyme is an area known as the active site. This has a very specific shape, so only one substrate or type of substrate (with the right shape) will fit into the gap. The enzyme and substrate then slot together to form a complex, as a key fits into a lock. In this complex, the substrate is enabled to react at a lower activation energy. This may be due to bonds within it being deformed and stressed in the complex, so making them more likely to react. Once the reaction has been catalysed, the products are no longer the right shape to stay in the active site and the complex breaks up, releasing the products and freeing the enzyme for further catalytic action, (some enzymes break a substrate down into two or more products – a catabolic reaction, others bond two or more substrates together to form one product – an anabolic reaction)

Induced Fit Theory - However, more advanced biological techniques have shown that the active sites of enzymes are not the rigid shapes once thought. In the induced fit theory, the active site is thought of as having a distinctive but flexible shape. Once the substrate enters the active site, the shape of that site is modified around it to form the complex. Once the products have left the complex, the enzyme reverts back to its original shape until another substrate molecule binds. The latter process is sometimes termed dynamic recognition.

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Diagram of induced fit

Enzymes are sensitive to changes in their environment. Changes in temperature or pH can cause changes in the shape of the molecule, which affects its activity. Changes in the concentration of both the enzyme and its substrate will also affect the rate of an enzyme-catalysed reaction.

As I am concentrating on the relationship between temperature and the rate of activity of dehydrogenase enzymes in different yeast cells, I shall explain why this occurs. A rise in temperature increases the rate of reaction until the enzyme denatures. As the temperature increases, the kinetic energy of the substrate and the enzyme molecules increase, so they move faster and with more force, so there is a greater chance of successful collisions, leading to a greater rate of reaction (collision theory). At the optimum temperature of an enzyme (normally 45  o C), the reaction rate is at a maximum. But, if the temperature reaches too high a point (i.e. over optimum temperature), most enzymes lose their tertiary structure and they start to denature. The enzymes will have gained so much kinetic energy, they start to change shape themselves so that the substrates no longer fit into the active sites. They change shape because the bonding becomes irreversibly changed so the active site is permanently damaged. At this point, the rate of reaction rapidly starts to decrease. Even though at very high temperatures, the number of collisions is extremely high, without the active sites, no products can be formed.

The effect of temperature on the rate of reaction can be expressed as the temperature coefficient, Q 10.  For every 10  o C rise in temperature, the rate of reaction will have doubled:

Q 10 = Rate of reaction at (x +10) o C

                Rate of reaction at x o C

At decreasing temperatures (lower than the optimum), the rate of reaction decreases because of reduced enzyme/substrate collisions.

There are exceptions to these statements though. Thermophile enzymes have an optimum above 45 o C, mesophile enzymes have an optimum between 20  o C and 45  o C and psychrophile enzymes are still efficient below 20  o C.

Graph of rate of reaction against temperature with explanation.

Prediction: For the practical I will be carrying out, I predict, that as I increase the temperature up to the optimum point (around 45  o C), the rate of reaction will increase in direct proportion to the temperature. So the colour change of the solution will steadily take a shorter time to occur. When the temperature hits the optimum point, the rate of reaction will be at its maximum. This will show the fastest colour change. As the temperature continues to rise above the optimum, the enzymes will start to denature and it will take longer for a colour change to occur. The rate of reaction will rapidly decrease as the enzymes are given so much kinetic energy, they start to change shape so the substrates don’t fit in exactly anymore. They change shape because the bonding becomes irreversibly changed so the active site is permanently damaged. The experiment will eventually reach a temperature where the enzymes will be totally denatured and will have completely changed shape, so the rate of reaction will be zero.

Outline method: this experiment is designed to show the relationship between temperature and the rate of activity of dehydrogenase enzymes in different yeast cells. triphenyl tetrazolium chloride (ttc) is an artificial hydrogen acceptor, or redox indicator. when oxidised, ttc is colourless, but when it is reduced, ttc will form red, insoluble compounds called formazans. this colour change shows the presence of active dehydrogenase enzymes in yeast cells. the temperature of the ttc solution and yeast suspension will affect the rate at which this colour change occurs, which in turn will show how the activity of dehydrogenase enzymes in different yeast cells changes..

Set up of apparatus

Proposed Experimental Method:

• Set up a water bath at 10  o C, adding crushed ice if necessary.
• Put 10cm 3 using a pipette of ‘bakers’ yeast suspension into a test tube labelled A.
• Put 1cm 3  using a pipette of TTC solution into another test tube labelled B.  
• Place both test tubes A and B into the water bath using tongs, and leave for several minutes to reach the temperature of the water bath. Check the temperature by using a thermometer.
• By using tongs mix together the two solutions by placing the contents of test tube A into test tube B (using tongs) and return to the water bath.
• Start the stop clock immediately.
• Observe the test tube carefully and record the time taken (in a table) of the duration it took for a colour change to occur. Stop the stop clock immediately.
• Repeat steps 1 – 7 using different temperatures for the water bath (in point 1.): 20  o C, 30  o C, 40  o C and 50  o C.
• Repeat steps 1 – 8 except in point 2. use ‘brewers’ yeast instead of ‘bakers’ yeast

Repeat the experiment 3 times for each yeast, ‘bakers’ and ‘brewers’, over the 5 different temperatures.

• Take averages of all the data and record in appropriate column of table.
• Plot a graph of temperature,  o C, (x axis) against rate of reaction, 1 / time , (y axis) using the average values.

Variable Identification and Control:

• Temperature  – I am going to vary the temperature of this experiment using set values of 10  o C, 20  o C, 30  o C, 40  o C and 50  o C. To get these set temperature values, I will change the temperature of the water bath by either heating it up or adding crushed ice. This will help me to draw a conclusion of the relationship between temperature and the rate of activity of dehydrogenase enzymes in different yeast cells.
• Substrate Concentration  – The substrate concentration must remain the same throughout the whole experiment. At high substrate levels the number of collisions increases, as the reacting particles are closer together and more active sites are used up. At low substrate levels, the active sites are not used up as there are not enough substrate molecules to occupy them all. Increasing the concentration of the substrate increases the rate of reaction until the enzyme concentration limits the rate of reaction. I can control this by using the same concentration of TTC from the same bottle of solution.
• Enzyme Concentration  - The enzyme concentration must remain the same throughout the whole experiment. As the concentration of the enzyme increases, the number of active sites increases, so there are more sites for substrate molecules to combine with. As the concentration of the enzyme decreases, the number of active sites decreases, so there are more uncombined substrate molecule. Increasing the enzyme concentration increases the rate of reaction until the substrate concentration limits the rate of reaction. I can control this by using the same concentration of actively respiring yeast suspension from the same bottle of solution.
• pH  – Each enzyme has its own characteristic pH at which the reaction will proceed at the fastest rate. Above and below the optimum value, the reaction will proceed more slowly so I must keep the pH value constant throught the experiment. At extreme pH values the enzyme will become denatured, and the shape of the protein molecules are altered as the hydrogen bonds and sulphur bridges are broken or formed. I can control this by using the same pH of enzyme and substrate from the same bottles of their solutions, and double check by using universal indicator paper.
• Volume  – I must keep the volume of the yeast suspension and the TTC solution the same. If I put a large volume of TTC into a test tube and react it with the yeast at a set temperature, the rate of this reaction would be faster than if I put a small volume of TTC into a test tube and reacted it with the same yeast at the same temperature. To control this I will measure the volume (in cm 3 ) using a measuring cylinder of each solution.

Reliable results:

Range of temperatures ( o C):                         10  o C, 20  o C, 30  o C, 40  o C and 50  o C.                         (Measured with thermometer)

Types of yeast:                                 ‘Bakers’ and ‘Brewers’.

Concentration of enzyme and substrate (mol):Kept the same throughout. Recorded at each set temperature.

Volume of enzyme and substrate (cm 3 ):         Kept the same throughout. Recorded at each set temperature. (Measured with measuring cylinder)

pH of enzyme and substrate:         Kept the same throughout. Recorded at each set temperature.

Number of repetitions of each yeast at

the set temperature:                                3

Risk Assessment and Ethical Considerations:

See ‘Material Data Safety Sheet’ for TTC.

Always wear safety goggles to prevent solutions splashing into eyes.

Document Details

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• Subject Science

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Mitochondrial copper overload promotes renal fibrosis via inhibiting pyruvate dehydrogenase activity

• Original Article
• Open access
• Published: 09 August 2024
• Volume 81 , article number  340 , ( 2024 )

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• Saiya Zhu 1   na1 ,
• Yangyang Niu 1   na1 ,
• Wenqian Zhou 1 ,
• Yuqing Liu 1 ,
• Jing Liu 1 ,
• Limin Lu 2 &
• Chen Yu   ORCID: orcid.org/0000-0001-7169-276X 1  

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Copper is a trace element essential for numerous biological activities, whereas the mitochondria serve as both major sites of intracellular copper utilization and copper reservoir. Here, we investigated the impact of mitochondrial copper overload on the tricarboxylic acid cycle, renal senescence and fibrosis. We found that copper ion levels are significantly elevated in the mitochondria in fibrotic kidney tissues, which are accompanied by reduced pyruvate dehydrogenase (PDH) activity, mitochondrial dysfunction, cellular senescence and renal fibrosis. Conversely, lowering mitochondrial copper levels effectively restore PDH enzyme activity, improve mitochondrial function, mitigate cellular senescence and renal fibrosis. Mechanically, we found that mitochondrial copper could bind directly to lipoylated dihydrolipoamide acetyltransferase (DLAT), the E2 component of the PDH complex, thereby changing the interaction between the subunits of lipoylated DLAT, inducing lipoylated DLAT protein dimerization, and ultimately inhibiting PDH enzyme activity. Collectively, our study indicates that mitochondrial copper overload could inhibit PDH activity, subsequently leading to mitochondrial dysfunction, cellular senescence and renal fibrosis. Reducing mitochondrial copper overload might therefore serve as a strategy to rescue renal fibrosis.

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Introduction

Kidney fibrosis is the common pathway and main histological manifestation underlying the progression of chronic kidney disease (CKD) to end-stage kidney disease [ 1 ]. The pathogenic mechanisms of renal fibrosis are diverse and involve a wide range of signalling pathways. Recent studies have shown that copper over-accumulation is associated with fibrosis in tissues such as the lung, the liver and the oral mucosa [ 2 , 3 , 4 ]. Copper is an essential micronutrient in living organisms and participates in many essential cellular processes [ 5 , 6 ]. In Wilson’s disease, the functional loss of ATPase copper transporting β (ATP7B) induces excessive copper accumulation in the liver, leading to liver cirrhosis, which can be ameliorated by copper chelator like penicillamine [ 7 ]. In our previous studies, we demonstrated that intracellular copper overload, driven by high expression of copper transporter 1 (CTR1), activated lysyl oxidase, enhanced matrix crosslinking and promoted renal fibrosis [ 8 ].

Mitochondria are crucial in eukaryotic cells due to their metabolic and biosynthetic functions. Mitochondria are not only vital intracellular copper reservoirs but are also the main copper utilizers [ 9 ]. Pyruvate dehydrogenase complex (PDHC) is a rate-limiting enzyme of glucose oxidation in mitochondrial matrix, converting pyruvate into acetyl-CoA and linking glycolysis to tricarboxylic acid (TCA) cycle [ 10 ]. Dihydrolipoamide acetyltransferase (DLAT), the E2 component of the PDHC, requires lipoylation for enzymatic function and exhibits a high affinity for copper within its lipoyl moiety [ 11 ]. A novel cell death pathway triggered by copper ionophore elesclomol, which is distinct from known death mechanisms and involves a 15–60 fold increase in intracellular copper levels, is discovered and coined “cuproptosis” by Tsvetkov et al. [ 12 ]. Our previous studies have shown that a mild accumulation of intracellular and mitochondrial copper, which is much lower than the fold increase mentioned in “cuproptosis”, induces cell damage and renal fibrosis [ 8 , 13 ]. However, the underlying mechanism requires further investigation.

In this study, we demonstrated that mitochondrial copper was elevated in fibrotic kidney tissues and TGF-β1-stimulated tubular epithelial cells. We further showed that mitochondrial copper might bind to the lipoyl moiety of lipoylated DLAT, leading to altered protein subunit interaction, lipoylated DLAT dimerization, and inhibited pyruvate dehydrogenase activity, thereby resulting in mitochondrial dysfunction and cellular senescence. Reducing mitochondrial copper levels by downregulating CTR1 expression or using a copper chelator were shown to restore mitochondrial function and attenuate renal fibrosis.

Materials and methods

Human kidney tissues.

Kidney biopsy tissues were acquired from the Department of Nephrology, Tongji Hospital, Shanghai, China. Control samples were obtained from patients with minimal change disease (MCD) without fibrosis, while fibrotic samples were collected from CKD patients with IgA nephropathy (IgAN) or diabetic nephropathy (DN). Masson trichrome staining, as previously reported [ 14 ], was used to evaluate the severity of renal fibrosis. The study was approved by the Human Subjects Committee of Tongji Hospital (No. K-W-2021-012), and written informed consent was obtained by all patients.

CTR1 +/− mice and the control mice (WT) were purchased from the Model Animal Research Center of Nanjing University and were routinely maintained on the C57BL/6 mice background. Polymerase chain reaction analysis of tail DNA was used to determine the targeted genes. All animals were housed in a room with controlled temperature (23 ± 2 ℃) and constant humidity (40–60%). All animals were provided with water and chow ad libitum. All animal experiments were approved by the Animal Ethics Committee of Tongji Hospital (No. 2020-DW-003).

For the unilateral renal ischaemia/reperfusion injury with contralateral nephrectomy (uIRIx) model, C57BL/6 male mice aged 6 to 8 weeks were used for the in vivo analysis. uIRIx models were established by clamping the left renal pedicle for 30 min using nontraumatic microvascular clamps at 37 °C. 7 days later, the mice were anaesthetized and the right kidney was removed. Twenty-eight days after the ischaemia, the mice ( n  = 5 per group) were killed, and the kidneys were harvested for analysis. Four groups of mice were prepared, namely, sham-operated WT mice (WT, sham), sham-operated CTR1 +/− mice (CTR1 +/− , sham), IRI 28d WT mice (WT, IRI 28d), and IRI 28d CTR1 +/− mice (CTR1 +/− , IRI 28d). For copper chelating experiments, copper chelating was achieved by daily oral treatment with TM (0.7 mg/day, Sigma-Aldrich) [ 15 , 16 ] for 28 days following the IRI operation. Mice were prepared into groups of sham-operated (Sham), sham-operated + TM-treated(Sham + TM), IRI 28d, and IRI 28d + TM-treated (IRI 28d + TM).

For the unilateral ureteral obstruction (UUO) model, C57BL/6 male mice aged 6 to 8 weeks were subjected to UUO operation as described previously [ 17 ]. Left ureter was exposed and ligated by surgical 5 − 0 silk. Mice ( n  = 4 per group) were sacrificed on day 14 after UUO surgery for biochemical and histological analyses.

For the folic acid (FA) model, C57BL/6 male mice aged 6 to 8 weeks were injected with 250 mg/kg of FA diluted in 300 mM NaHCO 3 intraperitoneally. Mice ( n  = 4 per group) were killed on day 28 after FA injection for biochemical and histological analyses.

Cell culture and treatment

Rat renal tubular epithelial cells (NRK-52E) were obtained from the Chinese Academy of Sciences (Shanghai, China). Cells were grown in DMEM (Gibco, #10,569,044) supplemented with 4% FBS and maintained at 37 °C in a 5% CO 2 air incubator. NRK-52E cells were treated with 5 ng/ml TGF-β1 (R&D Systems, USA), 10 µmol/L CuSO 4 (Sangon Biotech, Shanghai, China), with or without TM. A CTR1 shRNA plasmid was used to generate a lentivirus, which was then transfected to cells to downregulate CTR1.

Western blotting

NRK-52E cells and kidney tissues were lysed in RIPA buffer. Protein samples were separated by SDS–PAGE gels and transferred to polyvinylidene fluoride (PVDF) membrane. Subsequently, the membranes were blocked with 5% skim milk for 1 h and incubated with primary antibodies at 4 °C overnight, followed by incubation with a horseradish peroxidase-conjugated secondary antibody. The primary antibodies used were as follows: anti-GAPDH (Proteintech, #60004-1-AP), anti-Collagen I (Abcam, #ab-21,286), anti-α-SMA (Abcam, #ab-124,964), anti-CTR1 (Abcam, #ab-129,067), anti-p16 INK4A (Santa Cruz, #sc-1661), anti-γH2AX (Abcam, ab26350;), anti-DLAT (Proteintech, #13426-1-AP), anti-p21 (Proteintech, #10355-1-AP), anti-p53 (Cell Signaling Technology, #2524).

Reverse transcription and quantitative real-time PCR

RNA was extracted from cells and kidney tissues with TRIzol reagent (Invitrogen, Carlsbad, CA) as previously reported [ 18 ]. RNA was then transcribed into cDNA using a PrimeScript RT Reagent kit (Takara). Subsequently, cDNA served as a template for real-time PCR, which was conducted with SYBR Green PCR Master Mix (Roche, Mannheim, Germany). The primers used in this study targeted mouse GAPDH, Collagen I, CTR1 and α-SMA (Table  1 ), along with rat GAPDH and CTR1 (Table  2 ).

Renal histology and immunohistochemistry

Renal tissues were fixed in 4% neutral-buffered paraformaldehyde and paraffin-embedded according to a standard procedure. Kidney Sect. (4-µm thickness) were subjected to Masson trichrome staining, HE staining, or immunohistochemistry following the protocol [ 19 ]. For immunochemical staining, after the tissue sections were dewaxed and subjected to antigen retrieval, the sections were blocked with 3% hydrogen peroxidase for 10 min, followed by a 60-min incubation with 5% goat serum at room temperature. Next, the sections were incubated with a primary antibody against CTR1 (Abcam, #ab-129,067). Following incubation with an HRP-conjugated secondary antibody for 1 h at room temperature, a 3,3′-diaminobenzidine (DAB) kit (Vector, #SK4100) was used in accordance with the manufacturer’s instructions.

Transmission electron microscopy

To examine mitochondrial structure, transmission electron microscopy (TEM) was used. Kidney cortexes and NRK-52E cells were fixed in 2.5% glutaraldehyde/0.1 M phosphate buffer at 4 °C overnight. Subsequently, ultrathin renal sections were stained with uranyl acetate and lead citrate, then embedded in copper grid according to routine procedures. Mitochondria were photographed under an electron microscope (JEOL JEM-1010, Tokyo, Japan).

Isolation of mitochondria

Mitochondria from the kidney cortex and NRK-52E cells were isolated using commercial Mitochondria Isolation kit for Tissue (Abcam, #ab-110,168) and Mitochondria Isolation Kit for Cultured Cells (Abcam, #ab-110,170) following the manufacturer’s instructions respectively. The isolated mitochondria were then resuspended in mitochondrial storage buffer and BCA method was used to assess the protein concentration of the isolated mitochondria.

Copper content determination

Subcellular fractions of renal tissues and cells were treated with 65% nitric acid at 100 °C for 1 h, followed by diluted with ultrapure water. Copper levels were measured using inductively coupled plasma-mass spectrometry (ICP-MS) (NexION 300D, PerkinElmer, USA) as previously described [ 20 ]. Standard samples were also employed for quantification [ 21 ].

PDH enzyme activity

PDH activity in vitro experiment was measured using the PDH Enzyme Activity Assay Kit (Abcam, #ab-109,902) according to the manufacturer’s protocol. PDH activity in vivo experiment was assayed by PDH Enzyme Activity Assay Kit (Solarbio, # BC0385) as previously described [ 22 ].

Measurement of oxygen consumption rate

Seahorse XFe96 Extracellular Flux Analyzer (Agilent, CA) was used to detect oxygen consumption rate (OCR) according to the instructions provided by Seahorse Biosciences. Briefly, NRK-52E cells were seeded at a density of 6,000 cells per well in XF96 cell culture miniplates and incubated at 37 ℃ for 24 h. The cells were washed with XF assay medium supplemented with 2 mM glutamine, 1 mM pyruvate and 10 mM glucose, and equilibrated in 180 µL assay medium per well in a 37℃ incubator without CO 2 for 1 h. OCR was analyzed by injections of 1.5 µM oligomycin, 1µM carbonyl cyanide 4-(trifluoromethoxy) phenylhydrazone (FCCP), and 0.5 µM rotenone plus antimycin A to assess a mitochondrial stress test using the XF Extracellular Flux Analyzer. Following each assay, cell numbers were recounted for normalization of the OCR.

SA-β-gal activity assay

SA-β-gal staining was examined according to the manufacturer’s instructions (Beyotime, #C0602). Frozen kidney tissue sections were fixed with fixative solution at room temperature for 15 min and then washed three times with PBS. The fixed sections were incubated with a staining solution mix overnight at 37 ℃ without CO 2 . SA-β-Gal quantification was performed by Image J software [ 23 ].

ATP levels were detected using the ATP Assay Kit (Beyotime, #S0027) according to the manufacturer’s protocol. Briefly, cells and kidney tissues were homogenized in ATP lysis buffer and then centrifuged at 12,000 ×g and 4 ℃ for 5 min. 100 µL of ATP detection working solution was mixed with 20 µL of supernatant in 96-well plates. Luminance was then assayed and normalized to protein concentration.

Protein structure

The protein structure of DLAT was downloaded from the AlphaFold2 modelling results of the Uniprot website [ 24 ], where the UniProt number of DLAT is Q8BMF4. And lipoylation was performed on amino acids 131 and 258 of the DLAT. The pymol.edit module was used for initial structure establishment.

Molecular dynamics simulations (MDS)

To examine the interaction between lipoylation site and copper, wild-type and lipoylated DLAT protein structures underwent dynamics simulations. Gromacs 2019.6 was used as the dynamics simulation software [ 25 ] and GROMOS96 54a7 was used as the dynamic force field [ 26 ]. TIP3P water model was used along with NaCl and then one copper was added to construct control (DLAT-Cu 1+ (1)) and experimental (Lipoylated DLAT-Cu 1+ (1)) simulation system. Verlet, cg algorithms were used in the elastic simulation, with PME handling electrostatic interactions. Energy minimization used the steepest descent method, and the system was balanced using canonical (NVT) and isothermal-isobaric (NPT) ensembles. A 100 ns MD simulation at 300 K and normal pressure.

We took another simulation to examine the role of copper in lipoylation DLAT protein. TIP3P water model was used along with NaCl and then one copper was added to lipoylated DLAT system to construct one experimental simulation system (Lipoylated DLAT-Cu 1+ (1)). Lipoylated DLAT-Cu 1+ (0) were used as control group. The simulation, based on previously set parameters, involved a 10 ns MD simulation. Mirror the relaxed protein. SMD simulations on heavy atoms used a constraint force of 0.5 kcal/ (mol Å 2 ) to get initial coordinates for umbrella sampling along the unbinding pathway. A virtual spring constant of 5000 kcal/ (mol Å 2 ) was applied to the ligand’s centre of mass with a velocity of 0.01 Å/ps.

Determination of blood urea nitrogen and serum creatinine

The serum of the mice was isolated by centrifugation. Serum creatinine and blood urea nitrogen (BUN) levels were assessed using an automatic chemistry analyser according to the manufacturer’s manual. The levels of serum creatinine and BUN were expressed as µmol/L and mmol/L respectively.

Statistical methods

All results were presented as the mean ± SEM. Differences between two groups were analysed using Student’s t -test and one-way analysis of variance (ANOVA) was used for comparisons among multiple groups. Difference at p  < 0.05 was considered statistically significant. SPSS 21.0 statistical software (Chicago, IL) was used for statistical analyses.

Mitochondrial copper overload in fibrotic kidneys

Our previous data suggest that increased intracellular copper content contributes to renal fibrosis through lysyl oxidase mediated matrix crosslinking [ 9 ]. Here, we further investigated mitochondrial copper content in the fibrotic kidneys. We found that the mitochondrial copper levels but not the cytosolic levels were significantly elevated in the IRI 28d fibrotic kidneys, accompanied by accelerated cellular senescence, which can directly promote fibrogenesis [ 27 ] (Fig.  1 a-c). Similarly, mitochondrial copper levels were increased in the fibrotic kidneys of UUO-14d mice (Fig.  1 d-f) and FA-28d mice (Fig.  1 g-i), which were accompanied by elevated cellular senescence. In tubular epithelial cell stimulated by TGF-β1, we also found that the increased mitochondrial copper levels were associated with elevated expression of p16 INK4A and γH2AX (Fig.  1 j-k). These results indicated the tight association of mitochondrial copper overload with cellular senescence in the fibrotic kidneys.

Mitochondrial copper overload in fibrotic kidneys. ( a ) The cytosolic and mitochondrial fractions were isolated from kidneys of sham and IRI 28d kidneys. Copper content was detected in the cytosol and mitochondria in the kidneys of the IRI 28d model via ICP-MS. ( b ) Masson staining, HE staining and SA-β-gal staining in the kidneys of IRI 28d model. Original magnification, ×200. Bar = 50 μm. ( c ) Western blot analysis of Collagen I, α-SMA, p16 INK4A , γH2AX, p21 and p53 expression in sham and IRI 28d kidneys. ( d ) Copper content was detected in the cytosol and mitochondria in the kidneys of the UUO 14d model via ICP-MS. ( e ) Masson staining, HE staining and SA-β-gal staining in the kidneys of UUO 14d model. Original magnification, ×200. Bar = 50 μm. ( f ) Western blot showing the expression of Collagen I, α-SMA, p16 INK4A , γH2AX, p21 and p53. ( g ) The copper content was detected in the cytosol and mitochondria in the kidneys of the FA 28d model. ( h ) Masson staining, HE staining and SA-β-gal staining in the kidneys of FA 28d model. Original magnification, ×200. Bar = 50 μm. ( i ) Western blot analysis of Collagen I, α-SMA, p16 INK4A , γH2AX, p21 and p53 expression. ( j ) The copper content was detected in the mitochondria of NRK-52E cells after 48 h of TGF-β1 treatment. ( k ) Western blot analysis of Collagen I, α-SMA, p16 INK4A and γH2AX expression. Data are the mean ± SEM of at least 3 independent experiments. n  = 4 for in vivo experiments and n  = 3 for vitro experiments. * P  < 0.05, ** P  < 0.01, *** P  < 0.001, **** P  < 0.0001

CTR1 is significantly increased in renal fibrosis and contributes to the elevation of mitochondrial copper

CTR1 is the main regulator of copper influx in eukaryotes [ 28 ]. We then examined the CTR1 expression in various renal fibrosis models. Notably, renal CTR1 was significantly increased in the fibrotic kidneys of IRI 28d mice by immunohistochemistry analyses (Fig.  2 a). Consistent with this, western blot indicated that CTR1 was significantly upregulated in fibrotic renal tissues (Fig.  2 b). Furthermore, CTR1 was also highly expressed in the kidneys of UUO 14d and FA 28d fibrotic mice (Fig.  2 c-f), in NRK-52E cells treated with TGF-β1 (Fig.  2 g), and in fibrotic kidney tissues from diabetic nephropathy and IgA nephropathy patients (Fig.  2 h). To explore the effect of CTR1 on mitochondrial copper levels, we generated CTR1 heterozygous mice (CTR1 +/− ) as CTR1 homozygous deletion mice (CTR1 −/− ) were embryonic lethal and constructed a CTR1 knockdown cell lines in vitro. We found CTR1 knockdown markedly reduced the levels of the mitochondrial copper but not the cytosolic copper in kidneys of the IRI 28d mice and in NRK-52E cells treated with TGF-β1 (Fig.  2 i, j, Supplementary Figure S1 a-d). CTR1 thus was highly expressed in renal fibrosis and CTR1 deficiency alleviated mitochondrial copper accumulation.

CTR1 is highly expressed in renal fibrosis and induces mitochondrial copper overload. ( a ) Immunohistochemical staining of CTR1 in low magnification (left panel: magnification,×40; bar = 200 μm) and high magnification (right panel: magnification,×200; bar = 50 μm) in the kidneys of IRI 28d model. ( b ) Western blot analysis of CTR1 expression in sham and IRI 28d kidneys. ( c ) Immunohistochemical staining of CTR1 in low magnification (left panel: magnification,×40; bar = 200 μm) and high magnification (right panel: magnification, ×200; bar = 50 μm) in the kidneys of UUO 14d model. ( d ) Western blot analysis of CTR1 expression in sham and UUO 14d kidneys. ( e ) Immunohistochemical staining of CTR1 in low magnification (left panel: magnification,×40; bar = 200 μm) and high magnification (right panel: magnification,×200; bar = 50 μm) in the kidneys of FA 28d model. ( f ) Western blot analysis of CTR1 expression in sham and FA 28d kidneys. ( g ) Western blot analysis of CTR1 expression in NRK-52E cells treated with TGF-β1. ( h ) Immunohistochemical staining of CTR1, Masson staining and HE staining in the kidneys of patients with or without fibrosis. Original magnification, ×200. Bar = 50 μm. ( i ) The cytosolic and mitochondrial fractions were isolated from kidneys of wild-type control mice, wild-type mice that received IRI surgery, CTR1 +/− control mice and CTR1 +/− mice that received IRI surgery. Copper concentrations in the subcellular fractions were assessed via ICP-MS. ( j ) Copper content was detected in the mitochondria of control, CTR1-knockdown cell lines treated with TGF-β1( n  = 3). Data are the mean ± SEM of at least 3 independent experiments. n  = 5 for in vivo experiments and n  = 3 for vitro experiments. * P  < 0.05, ** P  < 0.01, *** P  < 0.001, **** P  < 0.0001. MCD, minimal change disease; DN, diabetic nephropathy; IgAN, IgA nephropathy

Reducing mitochondrial copper overload by knocking-down CTR1 mitigates cellular senescence and renal fibrosis

We then investigated the effects of reducing mitochondrial copper through inhibiting CTR1 on cellular senescence and renal fibrosis. Our results showed that the degree of renal fibrosis in the CTR1 +/− IRI 28d mice was markedly ameliorated, based on Masson trichrome staining, HE staining, mRNA and protein expression levels of Collagen I and α-SMA (Fig.  3 a-d). Moreover, with the knockdown of CTR1, the level of SA-β-gal activity and the expression of p16 INK4A , γH2AX, p21 and p53 in IRI 28d mice kidneys were significantly reduced (Fig.  3 e-f). We also found the knockdown of CTR1 mitigated p16 INK4A and γH2AX expression in NRK-52E cells treated with TGF-β1 (Fig.  3 g). Collectively, these results suggest that mitochondrial copper overload promotes cellular senescence and renal fibrosis.

Reducing mitochondrial copper overload by knockdown of CTR1 alleviates cellular senescence and renal fibrosis. ( a ) Masson and HE staining of renal sections among indicated groups. Original magnification, ×200. Bar = 50 μm. ( b , c ) RT–PCR showing the changes in Collagen I and α-SMA mRNA levels. ( d ) Western blot showing the expression of α-SMA and Collagen I among the groups as indicated. ( e )Representative staining micrographs showing SA-β-gal activity in different groups. Original magnification, ×200. Bar = 50 μm. ( f ) Western blot showing the expression of p16 INK4A , γH2AX, p21 and p53. ( g ) Western blot showing the expression of p16 INK4A and γH2AX. Data are the mean ± SEM of at least 3 independent experiments. n  = 5 for in vivo experiments and n  = 3 for vitro experiments. ** P  < 0.01, *** P  < 0.001, **** P  < 0.0001; # P  < 0.05 versus TGF-β1-treated cells or IRI 28d groups, # P  < 0.01, ### P  < 0.001

Reducing mitochondrial copper overload ameliorates mitochondrial dysfunction

To further examine the effect of mitochondrial copper overload on mitochondrial function, we assessed mitochondrial morphological changes following CTR1 knockdown. In the IRI 28d kidneys, we observed that following the knockdown of CTR1, mitochondrial swelling(with a loss of cristae)was improved (Fig.  4 a). Moreover, downregulation of CTR1 increased ATP generation (Fig.  4 b). In vitro in the NRK-52E cells, the deformation of mitochondria induced by TGF-β1 was also ameliorated after knockdown of CTR1 (Fig.  4 c). In the Seahorse tests, TGF-β1-induced alterations in the basal respiration, ATP-production dependent respiration, maximal respiration were all partially reversed in CTR1 knockdown cells (Fig.  4 d-e). ATP levels were also restored after knockdown of CTR1 (Fig.  4 f). Together, these data indicated that reducing mitochondrial copper overload attenuated TGF-β1-induced mitochondrial dysfunction.

Reducing mitochondrial copper overload alleviates mitochondrial dysfunction. ( a ) Representative images of mitochondria morphology detected by transmission electron microscopy in low magnification (upper panel: magnification, ×6800; bar = 1 μm) and high magnification (bottom panel, × 13,000; bar = 500 nm). ( b ) ATP content detection of kidney tissues. ( c ) Representative electron micrographs of mitochondria in NRK-52E cells of the indicated groups in low magnification (upper panel: magnification, ×7000; bar = 2 μm) and high magnification (bottom panel: magnification, ×15,000; bar = 1 μm). ( d , e ) Measurements of the OCR in NRK-52E cells among different groups. ( f ) ATP content detection of cultured cells. Data are the mean ± SEM of at least 3 independent experiments. n  = 5 for in vivo experiments and n  = 3 for vitro experiments. * P  < 0.05, ** P  < 0.01

Mitochondrial copper overload inhibits PDH enzyme activity via inducing the dimerization of lipoylated DLAT

We then performed a series of further experiments to explore the mechanism by which mitochondrial copper overload induces mitochondrial dysfunction and cellular senescence. Through molecular dynamics simulations that had been verified to be stable and viable (Supplementary Figure S2 ), we found that copper ions were in close proximity to the lipoylated DLAT protein but distant from the non-lipoylated DLAT protein (Fig.  5 a). According to the trajectory, copper ions bound to the lipoylated site of lipoylated DLAT with a tendency of forming stable postures (Fig.  5 b), enhancing the interactions between protein subunits (Fig.  5 c) and reducing resistance energy between protein subunits (from − 268.9 kal/mol to -65.42 kal/mol) (Fig.  5 d), thereby facilitating protein dimerization. Additionally, we examined lipoylated DLAT dimer in fibrotic kidneys and found lipoylated DLAT dimer was significantly increased in the renal tissues of IRI 28d mice. In CTR1 +/− IRI 28d mice, however, the levels of DLAT dimer were significantly decreased (Fig.  5 e). Since DLAT is a crucial component of the PDHC, we further explored whether the dimerization of lipoylated DLAT could inhibit PDH enzyme activity. On one hand, downregulation of CTR1 markedly ameliorated IRI-induced decrease in PDH enzyme activity (Fig.  5 f). On the other hand, in tubule cells treated with TGF-β1, downregulation of CTR1 led to a reduced level of lipoylated DLAT dimer and increased PDH enzyme activity (Fig.  5 g, h). These findings suggest that mitochondrial copper overload probably inhibits PDH enzyme activity in part through the dimerization of lipoylated DLAT.

Mitochondrial copper induces the dimerization of lipoylated DLAT and inhibits PDH enzyme activity.( a ) Molecular dynamics simulations of the distance between copper ions and lipoylated DLAT/non-lipoylated DLAT. ( b ) Molecular dynamics simulations showing the effect between copper and modification site of lipoylated DLAT. ( c ) Molecular dynamics simulations showing the forces between lipoylated DLAT protein subunits with/without copper. ( d ) Energy between lipoylated DLAT protein subunits with/without copper. ( e ) Western blot analysis of the dimer of lipoylated DLAT proteins among different groups in vivo. ( f ) PDH enzyme activity detection. ( g ) Western blot analysis of the dimer of lipoylated DLAT proteins among different groups in vitro. ( h ) PDH enzyme activity detection. Data are the mean ± SEM of at least 3 independent experiments. n  = 5 for in vivo experiments and n  = 3 for vitro experiments. * P  < 0.05, ** P  < 0.01, ** P  < 0.001

Treatment with copper chelator tetrathiomolybdate (TM) improves PDH enzyme activity and mitochondrial dysfunction

To further affirm the role of mitochondrial copper over-accumulation in mitochondrial function, we used a copper chelation agent, TM. In comparison with the untreated IRI 28d mice, TM treatment significantly restored the PDH enzyme activity in renal tissues of the IRI 28d mice (Fig.  6 a). In addition, mitochondrial swelling and fragmentation of cristae in the IRI 28 model were improved after TM treatment (Fig.  6 b). The reduction of ATP levels in the IRI 28d mice was mitigated after TM stimulation (Fig.  6 c). Furthermore, we found that the lipoylated DLAT dimer significantly increased in tubule cells stimulated with TGF-β1 and markedly decreased after TM treatment (Fig.  6 d). TM treatment restored TGF-β1-stimulated reduction in PDH enzyme activity (Fig.  6 e). TGF-β1-induced abnormal mitochondrial morphology, declined maximal respiration, and decreased ATP levels were improved as a consequence of TM treatment (Fig.  6 f-i). In line with above observations, copper levels were markedly increased in the mitochondria after CuSO 4 treatment (Supplementary Figure S3 a). Mitochondrial shrinkage and disorganized cristae in CuSO 4 -treated NRK-52E cells were mitigated following TM treatment (Supplementary Figure S3 b).

Copper chelator TM treatment improves PDH enzyme activity and mitochondrial dysfunction. ( a ) PDH enzyme activity detection. ( b ) Representative electron microscopy images of mitochondria in renal tubular cells among the indicated groups in low magnification (upper panel: magnification, ×2500; bar = 5 μm) and high magnification (bottom panel: magnification, ×15,000; bar = 1 μm). ( c ) ATP content detection. ( d ) Western blot analysis of the dimer of lipoylated DLAT proteins among different groups in vitro. ( e ) PDH enzyme activity detection. ( f ) Representative electron microscopy images of mitochondria in renal tubular cells among the indicated groups in low magnification (upper panel: magnification, ×7000; bar = 2 μm) and high magnification (bottom panel: magnification, ×15,000; bar = 1 μm). ( g , h ) Measurements of the OCR in NRK-52E cells among different groups. ( i ) ATP content detection. Data are the mean ± SEM of at least 3 independent experiments. n  = 4 for in vivo experiments and n  = 3 for vitro experiments. * P  < 0.05, ** P  < 0.01

TM treatment mitigates renal cellular senescence and fibrosis

We finally verified the effects of copper chelation on cellular senescence and renal fibrosis. As evidenced by histological examination, and Collagen I and α-SMA expression, TM treatment significantly reduced renal fibrosis (Fig.  7 a-d). In addition, copper chelation markedly attenuated the abnormal high level of SA-β-gal activity and the increased expression of p16 INK4A , γH2AX, p21 and p53 (Fig.  7 e, f). The IRI-induced upregulation of serum creatinine and BUN in the IRI 28d mice model were profoundly attenuated after TM treatment (Fig.  7 g, h). In cultured tubule epithelial cells, TM treatment also markedly reduced TGF-β1 or CuSO 4 -induced overexpression of p16 INK4A and γH2AX (Fig.  7 i, Supplementary Figure S3 c). Together, these results indicated that copper chelating could ameliorate cellular senescence and renal fibrosis.

TM treatment improves cellular senescence and renal fibrosis. IRI 28d or sham-operated mice were divided into four groups: (1) sham mice treated with saline; (2) sham mice treated with TM (0.7 mg/day); (3) IRI 28d mice treated with saline; and (4) IRI 28d mice treated with TM. ( a ) Masson and HE staining of renal sections from the indicated groups. ( b - d ) Western blot and QPCR showing the expression of α-SMA and Collagen I among the groups as indicated. ( e ) Representative staining micrographs showing SA-β-gal activity. ( f ) Western blot analyses of the expression of p16 INK4A and γH2AX in different groups. ( g ) Serum creatinine and ( h ) blood urea nitrogen levels in different groups. ( i ) Western blot analyses of the expression of p16 INK4A and γH2AX in different groups. Data are the mean ± SEM of at least 3 independent experiments. n  = 5 for in vivo experiments and n  = 3 for vitro experiments. ** P  < 0.01, *** P  < 0.001, **** P  < 0.0001; # P  < 0.05 versus TGF-β1-treated cells, ## P  < 0.01 versus IRI 28d groups, ### P  < 0.001

The present study demonstrates mitochondrial copper overload in experimental CKD models. The mitochondrial copper is shown to bind directly to lipoylated DLAT protein, induces lipoylated DLAT protein dimerization, and inhibits PDH enzyme activity, which leads to mitochondrial dysfunction, cellular senescence and renal fibrosis.

Tsvetkov et al. challenged traditional cell death dogmas and proposed “Cuproptosis” in Science , suggesting that pulse treatment with the copper ionophore elesclomol results in a 15–60 fold increase in intracellular copper level, triggering proteotoxic stress and ultimately leading to cell death [ 12 ]. Similar to the proposed cuproptosis, the effects of copper overload on renal cells as we have demonstrated are also mediated at least in part through the mitochondria. Distinct differences, however, exist between cuproptosis and copper overload-induced renal cellular senescence and fibrosis. Firstly, in our models, the increase in intracellular or mitochondrial copper is relatively small and probably not sufficient to cause cuproptosis. In our studies, a mild increase (approximately 1.5-2 fold) in copper ions level within renal tubular epithelial cells and mitochondria, which is significantly lower than the fold-increase of copper ions in cuproptosis, is sufficient to cause cell damage, renal senescence and fibrosis [ 8 , 13 ].

Cellular senescence permanently arrests cell proliferation, accompanied by the development of senescence-associated secretory phenotype (SASP) [ 29 ]. It is showed that decreased respiratory capacity, elevated ADP/ATP and AMP/ATP ratios, reduced mitochondrial membrane potential, increased production of oxygen free radicals, serves as both a cause and a consequence of cellular senescence [ 30 ]. In current study we pinpoint that mitochondrial copper overload could promote renal senescence and fibrosis in part by inhibiting mitochondrial PDH activity. Malthankar et al. reported previously that elevated levels of Cu 2+ , Mn 2+ and Zn 2+ can inhibit the activities of TCA cycle enzymes, such as citrate synthase and ketoglutarate dehydrogenase complex in neuronal cells and fungal cells [ 31 , 32 , 33 ]​. In line with this, in our study involving CKD mouse models and TGF-β1-stimulated NRK-52 cells, we observed that mitochondrial copper overload leads to a reduction in PDH activity within the TCA cycle, thereby impeding ATP generation, inducing cellular senescence and promoting renal fibrosis. Reducing mitochondrial copper content (using copper chelation with TM or CTR1 knockdown), on the other hand, rescued PDH enzyme activity, replenished ATP production, and ameliorated renal fibrosis. Wilson’s Disease (WD), a rare hereditary condition, is characterized by hepatic mitochondrial copper accumulation and lethal liver failure. Unlike the mechanisms in our research, in WD, hepatic copper accumulation induces mitochondrial membrane crosslinking and destruction. Using copper-cheating agents can reverse copper levels, preserve mitochondrial structural integrity and function, and thereby alleviate liver damage [ 34 ].

Our results also suggest a novel mechanism through which mitochondrial copper overload inhibits PDH enzyme activity. Transition metal ions, such as copper ions, are potent mediators for protein aggregation, as they can facilitate stable and rigid interactions between proteins on a small surface area [ 35 , 36 ]. Martina G et al. reported that copper ions mediate protein aggregation and are implicated in the progression of neurodegenerative diseases [ 37 ]. In Alzheimer’s disease, it has been discovered that copper ions bind to amyloid-β peptide (Aβ), inducing the formation of Aβ oligomers and ultimately exacerbating the neurotoxicity of Aβ-aggregation [ 38 ]. Pyruvate dehydrogenase complex (PDHC) is a mitochondrial matrix enzyme that catalyses the irreversible conversion of pyruvate generated by glycolysis into acetyl-CoA (coenzyme A), which can then be fuelled into TCA cycle [ 39 ]. Zhang et al. reported that PDH enzyme activity was reduced in the UUO model and in tubular epithelial cells incubated with TGF-β1 [ 40 ]. The PDHC consists of three different enzymes, including the E2 subunit, DLAT, which requires lipoylation (lipoic acid acylation) for its enzymatic function. The possibility of copper to bind to lipoylated DLAT directly is supported by the observed binding of copper to free lipoic acid, with a measured dissociation constant of 10 − 17 [ 11 ]. Our results, for the first time, demonstrated mitochondrial copper bound to lipoylated site of lipoylated DLAT, which further altered interactions between DLAT protein subunits and induced protein dimerization, thereby inhibiting the activity of pyruvate dehydrogenase and ultimately resulting in cellular senescence and renal fibrosis.

Regarding the uptake of copper ions by the mitochondria, preliminary studies suggested that copper chaperone COX17 could ferry cytosolic copper into mitochondria [ 41 , 42 ]. However, later experiments showed that COX17 functions only in the mitochondrial membrane space and cannot transport copper ions from the cytoplasm to mitochondria, contradicting this notion [ 43 , 44 ]. Mari et al. proposed that the tripeptide glutathione (GSH) may participate in copper entry into mitochondria, as it can be easily shuttled into mitochondria via porin channels in the mitochondrial outer membrane. Nevertheless, this idea was challenged by studies in GSH deficiency yeast, which showed an unchanged mitochondrial copper level [ 45 ]. SLC25A3, a member of the mitochondrial carrier family, transports the copper-ligand (CuL) complex across the inner membrane for storage in the matrix [ 46 ]. Dennis R et al. reported that the depletion of SLC25A3 results in a reduction of mitochondrial copper levels [ 47 ]. More studies suggested that a non-protein, anionic copper ligand (CuL) might mediate copper transport into mitochondria. Nonetheless, the specific molecular identity of the CuL remains unclear, and further investigations are warranted to validate this concept [ 6 , 45 ]. CTR1 is a major copper importer for intracellular copper uptake [ 48 ]. CTR1 was highly expressed in kidney and regulated by Smad3 signalling [ 8 ]. Since the specific mitochondrial copper importer is currently unknown, we used CTR1 knockdown transgenic mice, CTR1 knockdown renal tubular cell lines, and combined with copper chelators to investigate the underlying mechanisms by which mitochondrial copper overload induces cellular damage.

Collectively, our study demonstrates that mitochondrial copper might bind directly to lipoylated DLAT and induces its dimerization, thereby inhibiting PDH enzyme activity and resulting in mitochondrial dysfunction, cellular senescence and renal fibrosis. Reducing mitochondrial copper levels by inhibiting CTR1 or using copper chelation might be an innovate strategy to mitigate the progression of renal fibrosis (Fig.  8 ).

Schematic illustration of mitochondrial copper overload in renal fibrosis. This study highlights that mitochondrial copper overloaded in fibrotic kidneys will lead to the dimerization of lipoylated DLAT, inhibition of pyruvate dehydrogenase activity, mitochondrial dysfunction, and cellular senescence. On the other hand, reducing mitochondrial copper overload through knocking-down of CTR1 or copper chelator treatment might restore PDH enzyme activity and mitochondrial function

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Acknowledgements

This study is supported by the National Natural Science Foundation of China (No 82170696), SHDC12022104, ITJ(ZD)2201 and 23Y11908900.

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Saiya Zhu and Yangyang Niu contributed equally to this work.

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Department of Nephrology, Tongji Hospital, School of Medicine, Tongji University, Shanghai, 200092, China

Saiya Zhu, Yangyang Niu, Wenqian Zhou, Yuqing Liu, Jing Liu, Xi Liu & Chen Yu

Department of Physiology and Pathophysiology, School of Basic Medical Sciences, Shanghai Medical College, Fudan University, Yixueyuan Road, Shanghai, 200032, China

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Saiya Zhu conducted experiments, wrote and edited the manuscript. Yangyang Niu conducted experiments while revising the article and reviewed the manuscript. Wenqian Zhou and Yuqing Liu helped conduct animal experiments. Jing Liu and Xi Liu designed the figures. Chen Yu and Limin Lu designed and supervised the study.

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Zhu, S., Niu, Y., Zhou, W. et al. Mitochondrial copper overload promotes renal fibrosis via inhibiting pyruvate dehydrogenase activity. Cell. Mol. Life Sci. 81 , 340 (2024). https://doi.org/10.1007/s00018-024-05358-1

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Received : 13 February 2024

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Accepted : 09 July 2024

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DOI : https://doi.org/10.1007/s00018-024-05358-1

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