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Fructose 1,6-bisphosphate aldolase (more commonly referred to as aldolase and technically known as D-glyceraldehyde-3-phosphate lyase, EC is an ubiquitous glycolytic enzyme that catalyzes the reversible cleavage of fructose 1,6-bisphosphate (Fru 1,6-P2) to glyceraldehyde 3-phosphate (G3P) and dihydroxyacetone phosphate (DHAP).  The enzyme also catalyzes the cleavage of structurally related sugar phosphates including fructose 1-phosphate (Fru 1-P), an intermediate of fructose metabolism.  Comparative studies of aldolases derived from diverse sources have demonstrated the presence of two classes of Fru 1,6-P2 aldolase, as defined by the SCOPS database [2], with different catalytic and molecular properties [3]; Class I aldolases are found in animals, plants and green algae while class II aldolases are found in bacteria, yeasts and fungi.
The class I aldolases of animals and higher plants have been widely studied [5].  These enzymes are invariably tetrameric [6] and both amino acid-sequence [7] and nucleic acid sequences indicate that they are highly conserved and derived by divergent evolution from a common ancestral gene .  They have identical molecular weights and subunit structures, readily form mixed hybrids in vivo and in vitro [9], and catalyze the same overall reactions, albeit with different kinetics.
Three unique forms of class I aldolase have been detected in various tissues of vertebrate species , including man [12].  These three enzymes, aldolase A (isolated from muscle), aldolase B (isolated from liver) and aldolase C (isolated from brain) have all been purified to homogeneity from rabbit tissues [14] and have been extensively characterized.  It is clear that these isozymes are closely related.  However, it is also clear that each is a unique protein species.  The three forms are immunologically distinct [15], have different peptide maps, have distinguishable catalytic activities, have different chromosomal locations , and different gene sequences [22,23].
In humans, aberrant aldolase activity has been associated with several inborn errors of metabolism.  A genetic defect of human aldolase A is associated with nonspherocytic hemolytic anemia (NSHA) [25].  A deficiency in F1-P cleavage by aldolase B in the liver, kidney, and small intestine results in a disorder known as hereditary fructose intolerance (HFI) [28,29].

As previously stated, class II aldolases are commonly found in bacteria, yeast and fungi.  These aldolases, however, are not classified under the class I aldolase superfamily because of the lack of primary sequence similarity; rather, class II aldolases comprise another superfamily.  Members of the class I aldolase superfamily share a similar reaction mechanism and intermediates, that being the use of covalent catalysis using a Schiff base intermediate [32].  Class II aldolases utilize a catalytic metal cation in order to polarize the carbonyl oxygen of their substrate.  It is interesting to note that, although class I and class II aldolases differ in their reaction mechanisms and intermediates, both classes share an alternating (a/b)8 barrel fold which will be discussed further in the protein structure section to follow.


Aldolase is involved in glycolysis, gluconeogenesis, and fructose metabolism.  The catalytic mechanism of a class I aldolase utilizes a Schiff-base intermediate for the cleavage of Fru 1,6-P2 and Fru 1-P [34].  Class II aldolase requires Zn2+ in the active site [35].

Investigations of the vertebrate Class I aldolases have elucidated several important physiological roles, the most prominent of which is its central position in the glycolytic pathway.  Glucose phosphorylation and isomerization steps, which precede aldolase in the glycolytic pathway, convert glucose into Fru 1,6-P2.  Aldolase catalyzes the cleavage of this hexose into two triose phosphates, G3P and DHAP.  The reverse reaction, synthesis of Fru 1,6-P2 from these triose phosphates in the gluconeogenic pathway, is also catalyzed by aldolase.  Those tissues that have a functional gluconeogenic pathway possess a special aldolase isozyme, aldolase B, as well as phospho-fructose bisphosphatase and phosphogluco-phosphatase [37].

In addition, aldolase is essential for the metabolism of fructose [38-40].  This pathway begins by conversion of fructose into Fru 1-P, by fructokinase.  The fructose is subsequently cleaved by aldolase into glyceraldehyde and DHAP.  The glyceraldehyde produced is subsequently phosphorylated to G3P by triose kinase and these two triose-phosphates can then enter into the glycolytic or gluconeogenic pathways.  See table for the Kinetic Parameters.

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Protein Structure

Aldolase is a tetramer [6] and it is found in all eukaryotic cells [12].  Its 40,000 Mr monomeric unit has a tertiary structure characteristic of the TIM barrel class of proteins, consisting of eight alternating a-helices and parallel b-strands .  Enzymes with the aforementioned structure and whose substrates form a Schiff-base intermediate are members of a mechanistically diverse superfamily known as the Class I Aldolase Family [32].  As a member of this superfamily, aldolase has three conserved isozyme members, A, B, and C.  The greatest disparity amongst vertebrate aldolase isozymes typically lies in the carboxyl-terminal tail (See TABLE for exon comparisons of aldolase isozymes).  Even though, as previously stated, there exists similarity in the (a/b)8 barrel fold between class I and class II aldolase, there is no clear sequence similarity between the two classes.

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Structures without Ligands

The first low-resolution crystal structure of aldolase was elucidated at 5Å resolution for human aldolase A in 1986 [41].  This structure has been refined to 2 Å resolution [42,43], although there has been some criticism of the structure [44,45].  The crystal structure of the more extensively studied rabbit aldolase A enzyme was initially resolved to 5Å resolution by Sygusch et al. [46], later refined to 2.7 Å resolution [47], then to 2.3 Å, and more recently refined to 1.9 Å resolution [48].  All these previous structures have been substituted in the database by the 1.9 Å structure [1ADO].  Many of the class I aldolase structures have been determined except for the last 20 amino acids at the carboxyl terminus.  In the structures listed below an asterisk (*) denotes the inclusion of a model for this C-terminal region.  The number in parenthesis denotes the number of subunits in each structure file.

            Aldolase Class I Structures Available on Protein Data Bank.
                        Aldolase A

  • Human (2Å) [1ALD] *(1) Note: disputed structure
  • Human (2.1Å) [2ALD] *(4)
  • Rabbit (1.9Å) [1ADO] ****(4)
  • Rabbit (1.8Å) [1ZAH] ****(4)
  • Drosophila (1.9Å ) [1FBA] ****(4)

Aldolase B

Aldolase C

Non-vertebrate Structures

  • Archae (1.91Å) [1OJX] (10)
  • Leishmania (1.8Å) [1EPX] (4)
  • P. ginivalis (2.46Å) [2IQT] *(1)
  • Plasmodium(3Å) [1A5C] (2)
  • Trypanosome (1.9Å) [1F2J] (1)

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Ligand Bound Structures
The first liganded structure of aldolase was with dihydroxyacetone phosphate (DHAP) [48], the common product of the glycolytic metabolic pathway.  In 1999, two structures were published that revealed the potential dynamics of the substrate, fructose bisphosphate, by revealing two binding modes for the C6-phosphate of Fru 1,6-P2 [44,45].  For more information, see Biochemistry of Aldolase.

            Ligand Bound Aldolase I Structures Available on Protein Data Bank
Aldolase A

  • Human with Fru 1,6-P2 (2.8Å) [4ALD] *(1)
  • Rabbit with DHAP (1.9Å) [1ADO] see above
  • Rabbit-mutant with Fru 1,6-P2 (2.3Å) [6ALD] (4)
  • Rabbit with DHAP-covalent (2.65Å) [1J4E] (4)
  • Rabbit with Fru-1,6-P2-covalent (1.76Å) [1ZAI] ****(4)
  • Rabbit with mannitol 1,6-bisphosphate (1.89Å) [1ZAJ] ****(4)
  • Rabbit with tagatose 1,6-bisphosphate (1.89Å) [1ZAL] ****(4)
  • Rabbit with N-(4-chlorophenyl)-3-(phosphonooxy)naphthalene-2-carboxamide(2.05Å) [2OT1] (4)
  • Rabbit with Wiskott Aldrich Syndrome Protein (2.05Å)[2OT0] (4)

Aldolase B

  • Rabbit with Fru 1,6-P2, DHAP & 3-phosphoglycerol (2.1Å) [1FDJ] see above

Non-vertebrate Structures

  • Archae with DHAP (2.1Å) [1OK4] (10)
  • Archaewith Glycerol (2.4Å) [1OK6] (10)
  • Archae with Fru-1,6-P2-furanose (1.85Å) [1W8S] (10)
  • Archae with covalent Fru 1,6-P2 (1.93Å) [1W8R] (10)

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Structures of Mutant Forms
Site directed mutagenesis, a molecular biology technique used to mutate specified DNA bases to introduce alternative amino acids in specific sites of a protein, has been a very useful technique in studying the role and function of certain residues of the aldolase enzyme.  In addition, mutations are also commonly found in vivo.  In particular, mutations in the aldolase B gene are the common perpetrators of Hereditary Fructose Intolerance (HFI), which is a potentially lethal genetic disorder hindering sugar metabolism following fructose ingestion.  The most common mutation causes a proline substitution for alanine at position 149 of aldolase B .  This mutation leads to inadequate thermal stability, activity, and quaternary structure, which result in its inability to carry out normal function [49].  The structure of the HFI protein (A149P-substituted human Aldolase B) represents one of the few instances in pathology where the structure of the perpetrating entity has been resolved [50].

Mutant Aldolase Structures Available on Protein Data Bank
Aldolase A

  • Rabbit-K146A with Fru 1,6-P2 (2.3Å) [6ALD] see above
  • Rabbit-E187Q (2Å) [1EWG] ****(4)
  • Rabbit-E187A (2.2Å) [1EX5] ****(4)
  • Rabbit-K107M (2.46Å) [1EWD] ****(4)
  • Rabbit-K229M (2.6Å) [1EWE] ****(4)

Aldolase B

  • Human-A149P at 4C (3Å) [1XDL] (8)
  • Human-A149P at 18C (3Å) [1XDM] (8)

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Structures of Class II Aldolases
While the class II adolases have the same fold as the class I aldolases, they are not thought to have arrived by divergent evolution, rather are an example of convergent evolution of function [51].  They use different mechanisms.  The class II aldolases use a divalent metal at the active site.  Unless otherwise noted, each structure has a zinc ion at the active site.  The asterisk (*) indicates subunits with significant regions in the middle of the structure that are disorderd.

  • E. coli (1.67Å) [1DOS] (2)
  • E. coli (2.5Å) [1ZEN] *(1)
  • E. coli with phosphoglycohydroxamate (2Å) [1B57] **(2)
  • E. coli with Cd (2Å) [1GYN] *(1)
  • T. aquaticus with Co (2.3Å) [1RV8] *(4)
  • T. aquaticus with Yt and Co (2Å) [1RVG] *(4)
  • T. caldophilus with DHAP (2.2Å) [2FJK] *(4)

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Aldolase Isozymes

Aldolase A

Aldolase A is a glycolytic enzyme that catalyses the cleavage of fructose 1,6-bisphosphate to glycerol 3-phosphate and dihydroxyacetone phosphate (see Table of Kinetic Parameters).  A crucial role for aldolase A in physiology is demonstrated by human diseases that result from mutations in the aldolase A gene.  The only aldolase A mutations that have been discovered are relatively mild, leaving the enzyme activity relatively unchanged.  One mutation results in a thermolabile aldolase A, which has been shown to cause nonspherocytic hemolytic anemia (NSHA) [25].  This mutation causes an amino acid substitution in a region of intersubunit contact and destabilizes the oligomeric structure of the enzyme [52].  Little loss in activity was detected for these aldolase dimers.  Therefore, it appears that anemia is caused by a loss in stability of aldolase rather than innate activity.  Furthermore, this implies that null mutations in the gene for aldolase A, being the embryonic form of the enzyme in mammals [53], are likely lethal.  Other aldolase-A substitutions have similar effects [54-57].
Although there is a wide variety of aldolases, the majority of studies have focused on the Class I Fru-1,6-P2 aldolase of vertebrates, more particularly the enzyme isolated from rabbit muscle, aldolase A [58].  Class I Fru-1,6-P2 aldolase was first crystallized by Warburg and Christian [59] from rat muscle.  Taylor et al. [60] developed a simple procedure for the isolation of large amounts of aldolase from rabbit muscle that greatly aided in the further investigation of this enzyme, which ultimately led to rabbit aldolase A becoming the most extensively characterized Class I aldolase.  Rabbit aldolase A was the first aldolase for which an amino acid sequence was published [7].  Determination of the rabbit aldolase A mRNA sequence by cDNA cloning conclusively demonstrated that rabbit aldolase A had 363 amino acids and corrected several errors in the published amino acid sequence [61].
The tertiary and quaternary structures of rabbit and human aldolase A have been determined through studies utilizing x-ray crystallography.  Comparisons of the human and rabbit aldolase A structures reveal that they are nearly identical [44,47].  The carboxyl terminal region of the polypeptide is believed to be mobile, perhaps moving in and out of the active site, such that the electron density for these amino acids is often disordered (see protein structure.)
Aldolase A, despite being the most extensively studied isozyme of the aldolase family, has a catalytic mechanism that is not yet completely understood.  It is still unclear what roles are played by several key amino acid residues at the active site (see below).

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            Proposed Mechanism for Class I Fru-1,6-P2 Aldolases
An old proposal for the aldolase catalytic mechanism, which can be found in current biochemistry textbooks [37,62], was proposed in 1974 [7].  Numerous aspects of this mechanism have been discredited by analysis of the three dimensional structure of aldolase [42,43,47], comparison of the primary structures of the aldolase isozymes , and site-directed mutagenesis [63].  The use of site-directed mutagenesis and X-ray crystallography has afforded the elucidation of new mechanistic models.  The basic chemical steps for hexose cleavage includes binding, ring-opening, Schiff base formation, carbon-carbon bond cleavage, protonation of the product enamine, hydrolysis of the product Schiff base, and release of DHAP from the enzyme (see Figure).

Littlechild & Watson proposed a new model based primarily on analysis of the crystal structure of human aldolase A, which did not account for a good deal of functional data.  For instance, they suggested that the primary function of both Asp-33 and Lys-146 was catalysis of Schiff base formation, and that Tyr-363 is the residue involved in the acid/base chemistry of C3-C4 cleavage.  This is inconsistent with the relatively mild effect of mutations in the carboxyl-terminus on kcat [65].  Choi et al. (2001) proposed a new model that takes all of the available data into account [64].  The first component of this new proposal is a description of the interactions of active site amino acids and their ionization states in the free enzyme as derived from the crystal structure (Figure 1).  The model accounts for the kinetic data on site-directed mutagenesis of active site residues and structure of various enzyme-substrate complexes [32,45,48,66-70].  In addition, the effects of interactions on the pKa of the catalytic groups are proposed.

FIGURE 1:  THE ACTIVE SITE OF RABBIT ALDOLASE  Left)  Crystal structure viewed perpendicular to the b-barrel.  The image was created using RAS-MOL.  Lysines are red, aspartates are yellow, glutamates are green, arginines are purple and serines are cyan.  Solvent exposure to the active site is from the cleft to the left in this image.  Right)  A diagram of the same orientation of the active site as shown on the left.  The functional groups of the lysines, aspartates, glutamates, arginines, and serines are shown with the proposed ionization/protonation states before substrate binding as described in the text.

The ionization state of the free enzyme. The locations and ionization states of catalytic residues in the free enzyme are shown (Figure 1, right).  Asp-33 is deprotonated and its pKa is predicted to be quite low in the free enzyme because of the electrostatic interactions of this aspartate with Lys-107 and Lys-146.  The negative charge of Asp-33, in turn, stabilizes the positive charge of these lysines, increasing their pKa values.  Glu-187 also interacts with Lys-146.  This interaction serves to lower the pKa of Glu-187 such that it remains deprotonated in the free enzyme.  In addition, the negative charge of Glu-187 stabilizes the positive charge of Lys-146, also contributing to its increased pKa.  Lys-229, however, is deprotonated.  Its pKa is lowered because of its location deep in the active site in a relatively hydrophobic environment [45], its proximity to the positively charged Lys-146, an effect first described by Westheimer in 1938 [71] for acetoacetate decarboxylase.  The positive charges of Arg-303, Lys-107, and Lys-146 at the entry site for substrates form a binding site for these negatively charged molecules (Fru 1,6-P2, Fru 1-P, DHAP, G3P).

A step-by-step mechanism.  A proposal for the mechanism of catalysis of Class I Fru-1,6-P2 aldolases is described below.  The model is based on earlier steady-state measurement of catalytic intermediate [72-74], data using pre-steady-state kinetic measurements on site directed mutant enzymes [66-68,75] and structural evidence regarding the phosphate-sugar binding site [44,48].  This mechanism is largely the described in a 2001 publication [64].  A diagram of the steps outlined below is available.

(1) b-D-Fru 1,6-P2  furanose, the cyclic form of the sugar, is bound through interaction of the C1-phosphate with amide backbone groups of Ser-271, Gly-272,

Gly-302, and Arg-303, and through contact of the C6-phosphate with Arg-303 [45,69].

(2) The furanose ring is opened catalytically by the enzyme.  The site of this catalysis is unknown [69].  The open-chain hexose migrates to a new position in the active site such that the C6-phophate is now liganded to Lys-107 and Ser-38 [44,69].  Steric steering is accomplished by the methylene carbons of Arg-148, which is held securely in place by an ionic bond to Glu-189.  A key postulate of this mechanism is that upon interaction of Lys-107 with the C6-phosphate group the salt-bridge with Lys-107 to Asp-33 is lost.  This change, along with the presence of the negatively charged phosphates, increases the pKa of Asp-33.  The pKa of Glu-187 is also increased due to the proximity of the C1-phosphate.  As a result, of this increase in pKa, Glu-187 becomes protonated.

(3) There is nucleophilic attack by the e-amino group of Lys-229 on the si-face of the substrate's electron deficient carbonyl carbon [58].  There is an intra-molecular proton transfer from the lysine amino group to the oxyanion to form the 2(R)-carbinolamine [76].

(4) The protonated Glu-187 acts as the general acid [32].  Dehydration of the carbinolamine to the Schiff-base imine is catalyzed by the protonation of the carbinolamine hydroxyl group encouraging its leaving as water and the C2-Lysine bond collapses into the imine (Schiff base).  Water is released from the re-face of this incipient imine.  The resulting carboxylate of Glu-187 stabilizes the protonated imine/Schiff base.

 (5) The breaking of the C3-C4 bond of the substrate is initiated by the now-basic Asp-33 acting as a general base to pull off the C4-hydroxyl proton.  The proton from the C4-hydroxyl group is transferred to Asp-33.  As the electrons from the oxyanion of this C4-alcohol collapse into the aldehyde [64,77] glyceraldehyde 3-phosphate (G3P) is formed.  Electrons from C3-C4 bond create a C3-carbanion/enamine.  The reaction is favored by delocalization of these electrons using the cationic Schiff base acts acting as an electron sink.  The positively charged Lys-146 provides stabilization of the developing carbanion (negatively charged) intermediate [66,68].  In addition, there is tautomerization between the carbanion (ketimine) and enamine forms of the resulting three-carbon enzyme intermediate.  Help in the basicity of Asp-33 and the destabilization of the C3-C4 bond is achieved by the electron withdrawing ability of the Schiff base nitrogen, the stabilization of carbanion intermediate by the positive charge of Lys-146, and the resonance stabilization of products through the Schiff base.

(6) The first reaction product, G3P, is released.  As G3P leaves there is an immediate reduction in the pKa of Asp-33 as the salt-bridge with Lys-107 reforms due to the loss of the former C6-phosphate of G3P.  This model has difficulty explaining the pKa-shift of Asp-33 when Fru 1-P is the substrate, which does not bind to Lys-107.  The mechanistic differences between Fru 1,6-P2 and Fru 1-P have not been studied in much depth and many details remain unknown.

 (7) The carbanion/enamine intermediate that remains covalently bound to the enzyme at this mid-point in the catalytic cycle is protonated stereo-specifically by this now acidic Asp-33.  The residue that protonates the carbanion has been long sought.  It is not clear that the Asp-33 is indeed that player; there is evidence that this step is catalyzed, or at least promoted, by residue(s) in the carboxyl-terminus [65,72,78].  There is a conserved Tyr-363 which is found in all class I aldolases.  In this mechanism, the role of Tyr-363 is undefined and remains unknown.

(8) Hydrolysis of the imine/Schiff base with the product, DHAP, is catalyzed by Glu-187 in the reverse of the steps for its formation.  The pKa of Glu-187 is increased by the presence of the carboxylate anion of Asp-33.  Glu-187 accepts a proton from a water molecule promoting attack by the water oxygen at the carbon on the cationic imine.  There is evidence from 18O-exchange that this water may be structured and remain near the Schiff base during the catalytic cycle [79].  The mechanisms of other aldolases have invoked other structural waters [80].  The proton is transferred to the nitrogen of the resulting carbinolamine.

(9) The C-N bond of the carbinolamine is cleaved as the hydroxyl proton is accepted by Glu-187.

(10) As DHAP is released, Glu-187 loses its proton due to a reduction in its pKa because of the absence of the C1-phosphate.  This product release is the slow step in the overall reaction for aldolase A [66,73] and likely involves a conformational change of the carboxyl-terminus [65].

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Aldolase B

The aldolase B isozyme possesses a crucial role in fructose metabolism and gluconeogenesis.  Mutations in the human aldolase B gene, which result in diminished aldolase B activity, are the cause of an autosomal recessive disease, hereditary fructose intolerance (HFI) [85,86].  HFI leads to excessive levels of fructose 1-phosphate (Fru 1-P) in tissues expressing aldolase B.  Several hazardous metabolic effects result in liver and kidney damage [87,88].  The symptoms of the disorder (abdominal pain, vomiting and hypoglycemia) result from the ingestion of fructose or similar sugars that are metabolized through Fru 1-P [29].  This disease poses the greatest risk to the patient during weaning while in infancy, with potential lethality.  Growth retardation, renal tubular dysfunction, liver failure, coma and death may occur if the disease remains undiagnosed and the patient continues to ingest fructose [89-91].
The liver isozyme, aldolase B, has the greatest catalytic efficiency (kcat/Km) for both Fru-1-P cleavage and fructose-1,6-bisphosphate (Fru 1,6-P2) synthesis (see Table of kinetic parameters).  The ratio of the catalytic efficiencies when catalyzing reactions involved in fructose metabolism relative to those involved in glycolysis (Fru-1-P cleavage/ Fru-1,6-P2 cleavage) shows that aldolase B has a 15 to 30-fold greater efficiency over other isozymes.  Furthermore, the catalytic efficiency ratio of gluconeogenesis to glycolysis (synthesis/cleavage) shows aldolase B is also the most efficient isozyme having an eighteen-fold higher ratio of specificity constants over aldolase A and a three-fold increase over aldolase C.  It appears that aldolase B is well suited for its dual roles of gluconeogenesis and the metabolism of fructose in the liver.  The X-ray crystallographic structure of human aldolase B is very similar to aldolase A [44].  Furthermore, the three-dimensional structure of a pathological form of human aldolase B, which is found in most HFI patients, has been determined [50].
The expression of aldolase B is restricted to the liver, kidney cortex and small intestine.  However, the control mechanisms of aldolase B expression are not yet known.  Glucocorticoids are released upon hypoglycemic stress; these hormones control the transcription of other genes for gluconeogenesis, such as phosphoenolpyruvate carboxykinase (PEPCK) [92].  The transcription of the aldolase B gene is itself stimulated by glucose and insulin [93].  Furthermore, the mRNA levels of aldolase B have been shown to increase in experimental studies where rats are fed fructose [94].  The promoter in aldolase B has several liver specific cis-elements (HNF-1 and HNF-2 [95]) both within 200 base pairs upstream of the 5'-start of transcription [96,97] and in an element in the first intron [98-101].

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Aldolase C
The third vertebrate aldolase isozyme, aldolase C, is found in the brain and other tissues of neurological origin [12,14].  This enzyme has catalytic properties intermediate to those of aldolases A and B (see Table of kinetic parameters)[102].  Although the specific physiological role of this enzyme is unknown, it has been suggested that aldolase C, like other isozymes such as hexokinase I and lactate dehydrogenase-H, is found in tissues of constant metabolic activity.  Recent evidence indicates the occurrence of aldolase C in some tissues that can substitute for aldolase B in fructose metabolism [103].
Comparisons of amino acid and nucleotide sequences have shown that aldolase A and C are more closely related to each other than either is to aldolase B.  In contrast, the kinetic differences between aldolases A and C are not sufficiently distinct to explain why both isozymes are expressed in the brain.  This phenomenon could be explained by the subtle differences found during glycolysis in the neural cells that express aldolase C.  Alternatively, minor structural differences could be essential for supramolecular associations particular to each isozyme, i.e. aldolase A's actin-binding property [104] or aldolase C's S100 protein-binding property [105](see moonlighting).  Although expression of aldolase C has been reported in Xenopus ovary [106], it is not clear what role the isozyme has in this tissue.  Aldolase C is distributed heterogeneously in the mammalian brain [107] associated with the brain S-100 protein [105].  Furthermore, aldolase C has been identified as zebrin II, an intracellular antigen associated with sagittal organization of the mammalian cerebellum which structures itself in bands with sets of Purkinje cells giving a striated appearance .  In addition, aldolase C has been identified as an inositol trisphosphate binding protein in tracheal smooth muscle [108,109], however, the implication of this association is unclear.
Comparative analysis done in this study reveals that four of the five C-isozyme specific residues; Arg-314, Thr-324, Glu-332, and Gly-350 are located in the carboxyl-terminal region of the enzyme [110,111].  This region of the aldolase is thought to be involved in the determination of isozyme specific function [65].  The three dimensional structure of aldolase C [112] is similar to that of aldolases A and B [47,113].  All three isozymes have nearly identical active sites [112] and all of the isozyme specific residues are located outside of the active site [111].

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The Expression and Physiological Roles of Aldolase Isozymes
Three isozymes of aldolase have been identified in vertebrates .  These isozymes are encoded by three distinct, but related, genes that are derived from a single ancestral gene .  The three aldolases are expressed in a tissue specific manner.  In mammalian embryonic tissues, aldolase A is the primary isozyme, while in birds and amphibians, aldolase C serves as the primary embryonic form .  In the adult organism, aldolase A is found in most tissues, with especially high concentrations found in the muscle tissue [12].  Aldolase B is found mainly in the liver, the kidney cortex and, to a lesser extent, in and small intestinal mucosa [12].  Aldolase C is found in the brain, nervous tissue [14], and smooth muscle [109,114].
Though the kinetic parameters are different amongst the isozymes (see Table of kinetic parameters), they are consistent with their physiological roles.  Aldolase A’s turnover number is fifty-fold higher toward the glycolytic substrate Fru 1,6-P2 than aldolase B and at least five-fold higher than that of C.  The turnover number of the B isozyme towards fructose metabolism substrate, Fru 1-P, is two to five-fold higher than the turnover numbers of aldolases A and C.  Furthermore, the Km of aldolase B toward the two trioses (G3P and DHAP) makes it kinetically well suited for its role in gluconeogenesis (See Table).  Aldolase B demonstrates an equal activity with Fru 1,6-P2 as it does with Fru 1-P.  Aldolase C exhibits catalytic properties intermediate to those of the A and B isozymes.  Not only can these three aldolases be distinguished by their catalytic characteristics; each isozyme demonstrates unique electrophoretic, chromatographic, and immunological properties [15].

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Additional Cell Roles: Molecular Moonlighting

Upon the completion of the human genome project, one of the most surprising findings was that the human genome consisted of a much smaller number of genes than had been previously predicted.  Once hypothesized to have in excess of 100,000 genes, the number of predicted open reading frames is now thought to be between 20,000 and 25,000 [115].  How do mammalian genomes achieve their complexity with so few genes?  One theory is that many gene products have more than one cellular function, so called “moonlighting” functions.  As long as the second function is beneficial to the cell and does not interfere with the primary function, it is advantageous for the cell to keep it (as reviewed by Jeffery [116]).  Not surprisingly, many glycolytic enzymes, which are evolutionarily very old, have been found to be moonlighting proteins, including aldolase.

In addition to its enzymatic roles in glycolysis, gluconeogenesis, and fructose metabolism, aldolase has been shown to participate in other non-metabolic processes.  Over the years of study on aldolases, ample reports of non-catalytic functions have been published.  In most cases, aldolase has been reported to bind proteins seemingly unrelated to glycolysis such as, F-actin [1,104,117,118].  This interaction has been demonstrated both in vitro and in vivo [104,119].

            Actin-binding protein – Aldolase.  One of the common threads of many of the aldolase moonlighting functions is the ability of aldolase to bind to the actin cytoskeleton.  It has been known for over 40 years that aldolase is capable of binding to actin filaments [1], although, as outlined below, the exact function(s) of this interaction is(are) unknown.  It is well documented that aldolase A plays a structural role in the F-actin cytoskeleton in vivo [1,117-120] and it has the ability to cross-link F-actin into a gel or form rafts in vitro [121-123]).  The physiological significance of this interaction has been corroborated by showing that actin-bound aldolase can be reversibly released in cells by addition of 2-deoxyglucose, an inhibitor of glycolysis [124].  As mentioned above, the biological reasons for this interaction remain unknown largely because knockout mutants of aldolase A are either inviable or have severe growth defects [125].  Moreover, if aldolase plays an adaptor role such as suggested for GLUT4 or SNX9 (see below), the quaternary structure may have an important role in several moonlighting functions of aldolase.  Aldolase tetramers can cross-link actin filaments in solution [104].  Dimers of aldolase can decorate actin filaments, but have no cross-linking ability.  Monomers of aldolase have little to no actin binding ability, indicating that the protein must be at least dimeric in order to bind actin (Tolan D, Allen K, Lehman W, Pirani A and Ritterson C, unpublished studies).  The aldolase/actin interaction is not just an in vitro effect; studies in 3T3 cells show aldolase binding to actin in vivo using fluorescently labeled aldolase [119].  Although many studies have predicted putative actin binding regions of the aldolase protein [104,118,120], there is yet no unequivocal evidence describing the exact aldolase/actin-binding site.

Text Box: Fig. 2. Interaction Diagram for aldolase moonlighting. Each numbered (red) line represents a report of a direct interaction. Those involved in endocytosis are boxed in blue. CCS; clathrin-coated structures. Citations:1 [1], 2 [4], 3 [8], 4 [10], 5 [11], 6 [13], 7 [16,17], 8 [18], 9 [21], 10 [24], 11 [26], 12 [27], 13 [30], 14 [31], 15 [33], 16 [36],Although the aldolase - F-actin interaction has been known for years, the likely far-reaching consequences of this interaction are poorly understood.  For example, there are several reports of aldolase acting as a “molecular adaptor” to the cytoskeleton.  The infection process of several types of protists, such as the malaria-causing Plasmodium involves such a molecular adaptor.  The parasite’s aldolase protein forms a bridge between its own F-actin and cell surface adhesin on the host cell allowing protist infection to occur [31].  Another extremely significant role in connection to the cytoskeleton is the interaction with the GLUT4 glucose transporter [21], which is key to the insulin response [126].  GLUT4 is the insulin-responsive glucose transporter in adipocytes and other insulin-responsive cells.  Upon stimulation with insulin, GLUT4 translocates from internal membranes to the plasma membrane thus allowing facilitated transport of glucose.  However, thexmechanism of insulin-stimulated GLUT4-vesicle sequestration and translocation remains unknown [126].  An interaction of aldolase A, F-actin, and the C-terminal domain of GLUT4 has been reported [21], which can be disrupted by glycolytic intermediates in 3T3–L1 adipocytes.  Aldolase has also been associated with several proteins involved in endocytosis; such as sorting nexin 9 (SNX9), dynamin2 (dyn2) [8], and nucleating Wiscott-Aldrich Syndrome protein (nWASp) [16,17].  More about these interactions is described below. 

Involvement with Endocytosis. Recent investigations have shown aldolase existing in a complex with the endocytosis proteins SNX9 and dyn2 [8].  Dyn2 is a ubiquitously expressed form of dynamin that is involved in AP2/clathrin-mediated endocytosis [127,128].  When SNX9/dyn2 are bound to aldolase in the cytosol, membrane localization of SNX9 is inhibited.  One hypothesis predicts that upon phosphorylation of SNX9, by an unknown kinase, SNX9/dyn2 are released from the aldolase complex and re-localized to the plasma membrane to fulfill their function in endocytosis [8].  In addition, actin also plays a role in clathrin-mediated endocytosis [13].  Disruption of actin dynamics inhibits formation of new clathrin-coated structures, and clathrin-coated vesicle internalization is inhibited when cells are treated with F-actin-disrupting reagents latrunculin A and jasplakinolide.  These interactions are highlighted in blue in Fig. 2.

  • Aldolase associates with the vacuolar H+-ATPase (V-ATPase) proton pump, which is found in the kidney as well as in some bone cells and is involved in maintaining the acidity of certain cellular compartments such as endosomes, lysosomes, Golgi-derived vesicles, central vacuoles in yeast and plants, and some clathrin-coated vesicles [129].  V-ATPase consists of two major subunits that are each made up of several smaller subunits.  Active pumping of protons via ATP hydrolysis requires the association of the two main subunits, V1 (the “head”) and V0 (the “stalk”).  Lu et al., have shown that aldolase can associate with three different subunits from V-ATPase (at least one from each main subunit).  Furthermore, aldolase is necessary for assembly of the two main subunits, as yeast strains missing aldolase could not assemble the V-ATPase pump [129,130].  Since the pump cannot function without both of its subunits, the group hypothesizes that aldolase is necessary for the activity of V-ATPase.


  • Aldolases A and C have been shown to play a role in mRNA stability.  It is thought that aldolase binding to a particular mRNA makes this mRNA less stable, since it competes with protein translation machinery such as the PolyA binding protein (PABP).  One example of this is the role aldolase plays in the stability of neurofilament (NF) mRNA.  Aldolase may also be responsible for helping to degrade the mRNA through possible nucleolytic activity [36,131].
  • Aldolase has been reported to bind to several other proteins, but the functional roles in these interactions are not yet characterized.  These proteins include:
    •  g-tubulin [33]
    • Heparin [27]
    • Band 3 in erythrocytes [30]
    • Phospholipase D2 [24]
    • Wiscott-Aldrich Syndrome protein (WASp) Buscaglia et al., 2006; St-Jean et al., 2007

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Inborn Errors in Metabolism

          Hereditary Fructose Intolerance
Hereditary fructose intolerance (HFI) is a common, with an incidence rate of about 1 per 20,000 persons [132], inborn error in metabolism that affects the proper digestion and degradation of a principle sugar, fructose.  This sugar has found an increased importance in the western diet [39,133,134] and has made life more difficult for those affected by HFI.  There has been much research done on HFI [29,135-138] since its discovery in 1958 [139,140].  For a more complete description of work on HFI, the Tolan Laboratory has a completely separate HFI web site where much more information about the disease, treatments, diagnosis, and implications can be found.

Contrary to HFI, an extremely common, non-hereditary disease that affects fructose tolerability is called fructose malabsorption.  This condition may affect up to 1 in 3 persons (  Fructose malabsorption manifests itself with problematic symptoms found in the gastrointestinal tract.

HFI has brought to light some major questions of basic metabolism.  These basic questions of biochemistry are described below.

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HFI Enigma Models
HFI patients do not have difficulty maintaining their blood-glucose levels in the absence of fructose [29].  This is surprising given that, by definition, they suffer from a catalytic deficiency in what is thought to be a key glucose-synthesizing enzyme, aldolase B.  Two hypotheses have been proposed to explain this enigma.  The first proposes that there is residual activity in the mutant aldolase Bs [141-144].  The second argues that there is compensating expression of the other aldolase isozymes [145,146].  There is evidence for both hypotheses.  There are many reports of HFI enzymes with residual activity [141,143,144], as well as reports that residual aldolase activity is due to the persistence of the fetal form of aldolase, aldolase A [145], or the ectopic expression of the neural form, aldolase C [146].  According to the first model, the residual activity resides in mutant aldolase B enzymes in HFI patients.  The common missense substitution mutations, which are found in at least one allele in the majority of HFI patients , mostly, result in unstable proteins, some which have considerable residual activity [141,143,144].  In lieu of obtaining liver samples from an aldolase B-deficient patient, the second model can only be tested using an animal model for aldolase B-deficiency.  Such an animal model is not yet available.

Fructose Metabolism 
In HFI patients, only a small fraction of ingested fructose is excreted in the urine, which indicates that HFI patients eventually are able to metabolize nearly all of any ingested fructose.  While about 50% is assimilated by the liver and kidney, leading to pathology in those tissues due to the accumulation of Fru 1-P, the other half is metabolized at unknown sites in the body.  If these sites could be identified, then perhaps the tissues harboring these sites could be coaxed into accruing an increased burden of the ingested fructose and alleviate the pathology in the kidney and liver.  Other sites have also been implicated, such as the adipose [147] and the brain [103,148] tissues.  A test of this hypothesis could be done using an animal model for aldolase B-deficiency.  Again, such a model is not yet available.

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Non-spherocytic Hemolytic Anemia
Non-spherocytic hemolytic anemia (NSHA) is characteristic of deficiencies in glycolytic enzymes, which manifest themselves as disorders of erythrocytes [149].  Erythrocytes (red-blood cells) depend on glycolysis as their sole source of energy.  The loss of a glycolytic enzyme results in the inability to maintain cellular osmolarity via the ATP-powered ion pumps.  The most common of these glycolytic enzyme deficiencies are in pyruvate kinase [149,150]. 

There are rare cases of NSHA due to aldolase A deficiency [151].  One aspect of the aldolase-deficient NSHA is myopathy [55], likely due to the unknown need for an abundance of aldolase in muscle fibers.  Other reported manifestations are variable, including an association with mental retardation [150,151].  The mutations that cause aldolase A-deficient NSHA all appear to affect protein structure rather than activity [52,55,152].

The known mutations causing aldolase A deficiency are D128G [25,52], E206K [55], R303Op [54], C338Y [54], and G246S [56].  The obviously severe mutation that causes a stop codon at R303 was found as a heterozygous with the C338Y mutation.  The other mutations were found in individuals who were homozygous for these alleles, except the G246S allele, which was found in a compound heterozygous situation with E206K.

NSHA studies in the Tolan Laboratory have concentrated on the effects that these mutations have on the quaternary structure of aldolase.  In particular, we were the first to describe the causative effects of the first NSHA mutation, D128G and its affects on the quaternary structure where in the tetramer falls apart into an unstable dimer.  This has lead to further investigations on the role of the quaternary structure and its extraordinary stability in aldolase [153,154].

The subsequent work on NSHA has yielded a description of new mutations, E206K, which affects the other dimer interface in the aldolase tetramer, and C338Y.  Studies on the later are ongoing.

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Quaternary Structure of Proteins: The Aldolase Model

The study of the inborn error in metabolism, NSHA, has enlightened our general understanding of the role of quaternary structure of proteins.  Aldolase exists as a very stable hetero- or homo-tetramer with highly conserved subunit interactions [9,155,156].  We have characterized the first-described human mutation in the aldolase A gene that causes NSHA [25,52].  The substituted enzyme (D128G) has lost its native quaternary structure and this results in a loss of stability [52].  Several other substitutions showed similar results, including isosteric substitutions (D128N) and substitutions at other sites in the same subunit-subunit interface (Q125D).  The surprising corollary is that these dimeric aldolases are fully active.  Moreover, the study of the structure of the major mutant form of aldolase B that gives rise to another inborn error in metabolism, HFI, results in a dimeric enzyme [49].  The structure of this enzyme has been solved and showed that the same dimeric form as in the D128G form of aldolase A results [50].  This enzyme is very thermally unstable and only has at best 5-10% of normal activity [49].  A major conclusion from these studies shows that study of the defects arising from human mutations can reveal general principles of protein structure and function.

We extended this result and included substitutions at the other subunit-subunit interface.  These “double-mutant” forms of aldolase can exist as a monomer at certain temperatures, again retaining nearly full enzymatic activity [153].  In both mutant enzymes, the loss of quaternary structure is commensurate with a loss in thermal stability and provides evidence that thermal stability, for aldolase, is one major roles of the quaternary structure.  The interconversion of monomer and multimeric forms is very sensitive to temperature, as is the stability.  The aldolases that are defective in quaternary structure have been studied by analytical ultracentrifugation and x-ray crystallography.  Sedimentation equilibrium of native, dimeric, and monomeric forms of aldolase showed 106-fold decreases in dissociation constants [154], respectively.  The upshot of these studies is that aldolase has one of the strongest subunit-subunit associations known for any protein-protein interaction (10-28 M3).  In fact, other protein-folding studies have shown that the interface regions are so strong that they are among the last to unfold [157,158].  This has implications for alternative roles that may have been the driving force for evolution of this strong interface (see moonlighting functions).  X-ray structures of these mutant dimers, both aldolase A [159] and aldolase B [50], have not revealed large changes in subunit tertiary structure.

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Bionformatics and Metabolomics

The definition of exactly what comprises the field of bioinformatics is not always clear.  One definition would state that bioinformatics utilizes areas such as mathematics, statistics, and computer programming to solve problems in biochemistry and chemistry at the molecular level.  For the Tolan Laboratory, bioinformatics encompasses the use of genome-wide information for determining the physiology and biochemistry of sugar metabolism that could not be attained by previously available methods.  “Transcriptomics,” or gene-expression profiling on the whole-animal scale, or transcriptional profiling, is the determination of where and when genes are expressed.  Expansion of transcriptomics to a set of genes that are involved in a metabolic process is what we call physiolomics or metabolomics.  The later would include a determination of the presence or concentrations of small molecule metabolites.  What follows is a description of how each of these areas of bioinformatics is being utilized in the Tolan Laboratory.

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Gene Expression Profiling (Transcriptomics)
We have developed an accurate, simple, and fast algorithm that could predict the entire expression pattern/profile of any gene of interest within wild-type mice and humans.  This algorithm is called the virtual northern blot (VNB) [160].  Various other high-throughput methods of transcriptional profiling exist:  cDNA and oligonucleotide gene arrays, serial analysis of gene expression (SAGE), and quantitative reverse-transcription (qRT) PCR are now common, some being costly and labor-intensive.  However, the public domain expressed sequence tags (EST) databases (dbEST) provide an alternative for expression profiling that requires only an Internet connection [160-163].

EST’s are single-pass sequenced cDNAs representing expressed genes from a specific cell population or tissue [164].  They are 200-2000 nucleotides in length and they are derived from the partial sequencing of randomly primed, one, or both ends of a cDNA.  Since its inception in 1994, dbEST has grown exponentially.  Numerous EST mining algorithms [165-171] have been developed that take advantage of this tremendous and growing resource (>40 million sequences as of 2006; see Table of dbEST growth).  Expression profiling using the public dbEST is a useful and common method for exploring the transcriptome.  Nevertheless, gleaning reliable expression information from EST data has been challenging.  Manipulation of the mRNA population during construction of cDNA libraries and tissue dissection are factors that contribute to the sense that using dbEST for determination of gene expression profiles may be unreliable or problematic [160,172].  Resolution of these issues served as a driving force for the development of the VNB.  VNB addresses many of these issues and yields accurate and reliable expression profiles.

Both methods depend on dbEST; UNIGENE pre-clusters the data and tallies the EST-hits for each tissue; VNB generates the expression profile “on-the-fly” with the most recent version of the ever-growing dbEST, then tallies the EST-hits for each tissue.  However, for UNIGENE, if the EST cluster is incorrect for any gene, such as closely related paralogs, the expression data can be misleading [160].  Nevertheless, recently tools have been developed to accurately mine dbEST for quantitative expression profiling [160,167].  Methods for transcriptional profiling using dbEST [164] are available and use refined BLAST-based algorithms for assembling ESTs into contigs (clusters)(UniGene [173,174], CGAP [175], TIGR [176,177]).

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The concept of physiolomics, as utilized in the Tolan Lab, is depicted in the figure below.  The overlapping gene-expression profile gleaned from the profiles of individual genes involved in a pathway give candidate tissues where that metabolic pathway may operate.  This data can be validated to verify that this pathway indeed operates by various experimental methods.


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Fructose Metabolism

From the HFI enigma, it’s clear that fructose metabolism occurs in tissues other than the liver and kidney.  We have used a physiolomic approach using both UNIGENE and VNB to identify possible tissues for fructose metabolism [103,160].  This approach has been used to detect gene expression patterns as well as find novel genes [160,178].

Using either UNIGENE or VNB, the liver, kidney, and small intestine, showed expression of glut5, khk, aldoB and/or aldoC, and dak.  These genes encode the enzymes necessary for the metabolism of fructose via the fructose-1-phosphate pathway (see Figure).  In addition, expression of these genes was observed in the brain, breast, and lymphocytes.  Experimental validation of this pathway-specific bioinformatic approach is usually performed by immunohistochemistry and RNA in situ hybridization (RISH).  For the brain, expression of glut5 and khk was shown in neurons, in particular, in Purkinje cells [103].  This investigation introduces an approach for discovery of tissues and/or cell types engaged in fructose metabolism and will provide future targets for understand any adverse effects of high fructose intake.

Figure.  Fru-1-P and Fru-6-P pathways for fructose catabolism.  Pathways from fructose to pyruvate are depicted by arrows.  Plasma membrane is indicated by double line.  Transporters are all capital and critical enzymes are lowercase italics.  Fructose 1,6-bisphosphate (Fru 1,6-P2), 1,3 bisphosphoglycerate (1,3-bisPGA), 2-phosphoglycerate (2PGA), 3-phosphoglycerate (3PGA).


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From the gluconeogenesis enigma described above, it’s clear that gluconeogenesis may occur in tissues other than liver.  This hypothesis is partially based on evidence from HFI patients, where in the absence of fructose, HFI patients maintain a sufficient concentration of blood glucose even after an overnight fast [29].  Studies have confirmed that upon administration of dihydroxyacetone, a precursor in gluconeogenesis, a rapid rise in blood glucose is seen in HFI patients [179,180].  A similar physiolomic approach is being used to identify tissues other than liver that are capable of performing gluconeogenesis and participate in glucose homeostasis.

Other work supports this idea by showing that fructose-1,6-bisphosphatase and PEPCK, two gluconeogenic enzymes, have been found in human tissues where expression had not been previously known.  Immunohistochemistry showed these two enzymes are co-expressed not only in the liver and kidney, but also in the small intestine, stomach, adrenal gland, testis, and prostate [181].

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We are using mass spectroscopy and traditional biochemical approaches to discern the levels of metabolites of fructose and glucose metabolism in tissues identified by bioinformatics methods described above.


The Aldolase Super Family

Enzymes that utilize a Schiff-base intermediate formed with their substrates (see aldolase mechanism) and that share the same a/b barrel fold comprise a mechanistically diverse superfamily defined in the SCOPS database as the Class I aldolase family.  The family includes the “classical” aldolases fructose-1,6-(bis)phosphate (Fru-1,6-P2) aldolase, transaldolase (from the pentose-phosphate shunt), and 2-keto-3-deoxy-6-phospho-gluconate aldolase (from the Entner-Dudoroff pathway).  Moreover, the N-acetylneuraminidase lyase family [182] has been included in the Class I aldolase family based on similar Schiff-base chemistry and fold [32].  In addition, there are mechanistic similarities beyond the common use of a lysine for Schiff-base formation.  The structural and mechanistic correspondence comprises the use of a catalytic dyad, wherein a general acid/base residue (Glu, Tyr or His) involved in Schiff-base chemistry is stationed on beta strand 5 of the a/b barrel.  Site-directed mutagenesis and steady state and pre-steady state kinetics have demonstrated a role for this second member of this “catalytic dyad” [32,64] where it participates in the protonation of the carbinolamine intermediate and dehydration of the Schiff base.  Further mechanistic similarities are being investigated among members of this family to study the enzyme dynamics of “excursions” off the a/b barrel and the involvement of any dynamics on this region in substrate specificity.

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Molecular Biology of Aldolases

Modern molecular biology and the various genome projects have led to the deduction of the amino acid sequence of many genes and their corresponding proteins.  The ever-increasing database of primary protein sequences includes Class I aldolase isozymes from various species.  Analysis of aldolase primary sequences has provided insight into the extent of structural similarity that exists among aldolases.  Alignment of aldolase sequences from species ranging from plants to mammals reveals a high degree of similarity and conservation of amino acids at numerous positions.  There are 66 conserved amino acids, among the aldolases aligned (See alignment) [84,183].  Comparison of this information with the three dimensional structure of the enzyme reveals that 25 of these residues are within the b-strands that line the active site, 16 are in the corresponding a helices of the a/b structure, and 7 are at the subunit interfaces.  The carboxyl-terminal tyrosine residue is also conserved throughout Class I aldolases.

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Aldolase Gene Structure

Correspondingly, the genes for all vertebrate isozymes show the same structure.  The exons of all the known vertebrate aldolase genes are split by introns at precisely the same positions in the gene.  The 3'- and 5'-untranslated regions are of similar size, but very different in sequence.  The variation in intron size is considerable, ranging from 93 bp (D. melanogaster) to 416 bp (human B) in intron 7 and from 83 bp (human A) to 2937 bp (human B) in intron 8 [184-186].  You can view the generic vertebrate aldolase gene structure by clicking HERE.

A comparison of the overall DNA sequence similarity (Table) between the different aldolase isozymes reveals the greater degree of similarity (81%) that aldolase C has to aldolase A than to aldolase B (70%) [23].  This comparison revealed regions of higher and lower conservation between aldolase molecules.  Exons 4-7 in the aldolase A and C comparison are the most highly conserved whereas exons 3 and 9 have the least degree of similarity between isozymes.

While the structural genes are very similar among the aldolase isozymes, the promoter structures are quite distinct.  The aldolase A gene (aldoA) shows the most complexity of the three genes .  There are several alternative start sites for transcription and some alternative splicing for the 5’-untranslated regions [187].  Some of the promoters are used for the ubiquitous expression of aldoA in most tissues.  Other promoters are used in a more restrictive set of tissues, and yet a third promoter seems to be specific for the high levels of expression required for muscle cells.  Some of these promoters have canonical TATA boxes and others are TATA-less with the so-called, GC-rich promoter structure.  The aldolase B gene (aldoB) has a standard, well-described TATA-containing promoter, but in addition contains an enhancer sequence in the middle of the first intron [95-101].  The aldolase C gene (aldoC) has a CG-rich promoter without a TATA box [191,192].  The start of transcription is not well defined [191].

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Chromosomal Locations
The aldolase B gene was mapped to chromosome 9 using a rapid gene mapping system [193].  This “spot-blot” system uses a dual-laser sorter to identify and separate metaphase human chromosomes stained with either DIPI-chromomycin or Hoechst-chromomycin.  Chromosome panels were constructed from a normal cell line by sorting 22 chromosome fractions directly onto nitrocellulose filters.  Twelve labeled gene probes hybridized to the sorted chromosomal DNA fractions predicted by previous chromosome assignments.  The other genes have been mapped as described below.

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Human Aldolase Gene Chromosomal Locations
Spot-blot analysis of sorted chromosomes mapped human aldolase A to chromosome 16, aldolase C to chromosome 17, a pseudogene to chromosome 10, and the aldolase B gene to chromosome 9 (as mentioned above) [193].  Other reports using somatic cell hybrids or in situ hybridization and autoradiography have confirmed these loci [19,20,194].

The chromosomal locations of the HUMAN aldolase genes provide insight into their tissue-specific and developmentally regulated expression and evolution.  All loci are unlinked and located on two pairs of morphologically similar chromosomes consistent with tetraploidization during isozymic and vertebrate evolution (for more information on molecular evolution clink here).  Sequence comparisons of expressed and flanking regions support this conclusion .  These locations on similar chromosome pairs predict that the aldolase pseudogene arose when sequences from the aldolase A gene were inserted into the homologous aldolase location on chromosome 10 .  Lastly, there is evidence for another aldolase-related sequence on chromosome 3 [19,186].

Chromosomal Location of Human aldolase genes and pseudogenes
The chromosomal locations of the MOUSE aldolase genes show that they are similarly unlinked.  The Jackson Laboratory provides an excellent resource for information about the mouse genome.

Chromosomal Location of Mouse aldolase genes and pseudogenes      

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Developmental Expression of the Aldolase Genes
The tissue-specific expression of the vertebrate aldolase isozymes results from three developmental patterns of aldolase-isozyme gene expression.  In the first pattern, observed in mammalian muscle development, a relatively low level of aldolase A is expressed in the embryo from a specific promoter [187] until near birth when a dramatic increase occurs [53], which utilizes a distinct muscle-specific promoter [189,195].  This expression is the result of a switch in transcription between overlapping transcription units from a non-specific, or housekeeping promoter, to a muscle-specific promoter [189] (see molecular biology for a description of the promoters).
The second pattern involves a reciprocal switch in aldolase-isozyme gene expression.  This pattern has been observed in several tissues: developing mammalian liver, where aldolase B is expressed with a reciprocal disappearance of aldolase A [53]; in hepatocarcinogenesis, where this coordinated switch is reversed [196-198]; and in developing chicken muscle, where aldolase A is expressed with a reciprocal disappearance of aldolase C [199,200].
The third pattern has been observed in the developing mammalian brain where the expression of the embryonic aldolase A is joined by the coexpression of aldolase C [12,191].
The regulatory mechanisms that govern these patterns of developmental isozyme expression have been most extensively studied in mammalian muscle [12,53,189,195], where the first pattern arises from a transcriptional switch in cis.  The apparent coordinated switches found in the second pattern are less well characterized.  Chicken myoblast differentiation during early myogenesis in primary cultures offers a model for investigating the regulatory mechanisms of this second expression pattern.  In studies of chicken myogenesis [199] switch from aldolase-C isozyme expression to aldolase A isozyme expression occurred in an apparently coordinated fashion after the completion of myoblast fusion.  This reciprocal transition during chicken myogenesis is similar to the pattern observed in liver development, where transitions from A to B and from C to B occur in mammals and chickens, respectively [53].  We have recently shown that the switch is more complicated.  It is not coordinated at the level of the amount of steady-state mRNAs for aldolase C, which persists [200], and aldolase A, which is delayed relative to other muscle-specific genes [201].

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Historical References
For an extensive list of references documenting the discovery, cloning, DNA sequence determination, and characterization of the aldolase genes can be obtained by clicking here.

Molecular Evolution

The study of the relatedness of DNA and protein sequences is a powerful tool in understanding evolutionary processes.  Sequence comparisons in conjunction with a structural models can indicate conserved regions and their roles and relationships to each other, which cannot be found by comparison of primary sequence alone.  Functional genomics combined with structural and functional information can be extremely powerful in analysis of the important residues, understanding regions that are variable and finding regions that are specific to an isozyme type [183].

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Conservation of Gene Sequences
Having diverged from a common ancestor, the conservation of gene sequences for the aldolase isozymes is high.  Furthermore, this conservation indicates that they play fundamental roles amongst all species.  The level of genetic conservation usually gives insight as to how recent two species diverged from one another.  However, it is important to note that conservation is even seen between species, which have diverged very early on in the phylogenetic tree.  For aldolase, we see a high degree of conservation amongst all species that have been studied.  For example, in our studies, conservation the promoter elements of the aldolase B gene was shown to exist between rabbit, mammalian, and chicken systems [202].  In addition, the conservation of gene sequences can provide information for when homologous genes diverged.  From the similarity among the isozyme gene sequences, it has been proposed that aldolase B diverged from an A/C ancestor.  On the other hand, it has been proposed that aldolase B evolves faster than either aldolases A or C based on the assumption that all of the enzymes diverged from each other at the same point in time [203].  Our studies suggest that the non-synonymous sites (those that will not change the protein sequence) are evolving in a non-clock-like fashion and that aldolase B is accumulating non-synonymous changes at a significantly faster rate than aldolase A [183].

Times of divergence of aldolase isozymes.  An estimation of the divergence time of the vertebrate isozymes is instrumental in ascertaining the timing of duplication events that led to the evolution of these isozymes.  Since it has been shown that aldolase B accumulates non-synonymous changes at a faster rate than aldolase A, it is not valid to estimate the time of divergence of the aldolase A/C ancestor and aldolase B based on a constant rate of evolution for aldolase.  However, it is possible to calculate the time since the divergence of aldolases A and C since they are evolving at approximately the same rates.  These two isoforms diverged about 270 ± 70 million years ago [183].

Conservation of Aldolase Gene Structure.  The vertebrate aldolase genes that have been studied (human A, B, and C, rat A, B, and C, rabbit B, and chicken B) all share a common structure.  However, there is a large gap in the sampled taxa within vertebrate phylogeny.  In mammals and birds, the aldolase genes are composed of nine exons and eight introns, the junctions of which have been completely conserved [23,204] .  Outside of the vertebrate lineage, the fruit fly, Drosophila melanogaster, was the first aldolase gene structure investigated [185].  The D. melanogaster gene, however, has only four exons that correspond to the vertebrate exons 1, 2-7, 8 and 9 [185].  Vertebrate introns 1, 7 and 8 are present in D. melanogaster and the intron/exon junctions occur at precisely analogous bases.  The variation in intron size is considerable, ranging from 93 bp (D. melanogaster) to 416 bp (human B) in intron 7 and from 83 bp (human A) to 2937 bp (human B) in intron 8 [184-186].  The Drosophila aldolase genes have a unique structure, having a single locus that produces three mRNA transcripts [185,186,205].  The overlapping isozyme genes share a single promoter, a 5'-untranslated region, and two other protein coding exons.  The isozyme-specific carboxyl-terminal amino acids are encoded by one of three alternatively utilized fourth exons, by alternative splicing.  The three proteins differ in primary structure only at the carboxyl-terminal region encoded by the respective exon 4.  The enzymes (called a, b, and g) have been cloned, expressed in bacteria and characterized [206].  The three enzymes show similarity in optimal pH, thermal stability, and Km values for both Fru 1,6-P2 and Fru 1-P but have different kcat/Km values for the two sugar substrates.

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Conservation of Protein Sequences
Based on genetic conservation, it follows that proteins would also show conservation.  Because many species use many similar processes, be it metabolic or any other processes, it is logical that the proteins involved in these processes are conserved and, therefore, homologous (for a definition of this and related terms, and a description of its misuse, click here).  Protein conservation among the aldolases is high.  For a complete analysis, see Molecular Biology of Aldolases.  A phylogenetic tree based on parsimony [183] has been developed for the aldolases.

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Vertebrate Genome Evolution
A phylogenetic analysis will have significant impacts on our knowledge of the aldolase gene family, the fundamental mechanisms of vertebrate evolution, and the process of adaptive evolution.  It has been proposed that the animals that arose during the Cambrian explosion (500-420 mya) had nearly identical genomes (the "Cambrian pananimalia genome") and that all extant animal species evolved from these invertebrate, worm- and sponge-like ancestors [207].  How then was the vast variety of morphology and ecology of the vertebrates generated from essentially a single genome?  If the basic set of vertebrate genes were duplicated, this would create a source for variability on which selection could act.  The variation in vertebrates could then be explained by the evolution of novel functions in these duplicated genes/proteins in certain lineages and not in others.  The origin of multiple copies generated from a single (ancestral) gene may be explained by two mechanisms: tandem gene duplication or tetraploidization (genome duplication).

Tandem duplication may occur in several ways: (1) by unequal crossing over between homologous chromosomes in meiosis; (2) by unequal exchange between sister chromatids during mitosis; or (3) by regional redundant replication of DNA.  Logically then, tandem duplication leads to two copies of the gene in very tight linkage.

An alternative mechanism for gene duplication is by one or more duplications of the entire genome.  Tetraploidy may arise when two diploid (as opposed to the normal haploid) gametes fuse during fertilization and create a viable and fertile offspring.  Alternatively, a fertile triploid species with faulty meiosis mating with a normal diploid would also give rise to tetraploid offspring.  The now-tetraploid offspring, when mated to another tetraploid that arose through a similar mechanism, would give rise to a new tetraploid lineage.  The discovery of bisexual tetraploid and octaploid species of frog in South America [208] proves that this type of polyploid evolution is possible in animals.

In the Tolan Laboratory, we are gathering more evidence for, or against, the polyploidization model for the evolution of vertebrates [207,209-211].  This is an important question; evidence that tetraploidization has played a role in evolution of even one gene family requires that it was part of the evolution of most of the vertebrate genome.  The aldolase genes from many species are being isolated and characterized.  These species include; tunicate (a urochordate), amphioxus (a cephalochordate), and lamprey (a jawless fish).  Urochordates are the closest sister group to the cephalochordate/vertebrate clade, cephalochordates are the sister group to the vertebrate clade, and the jawless fish represent the most ancestral branch of the vertebrate tree [211] (see Figure).  The two isoforms, aldolase A and aldolase C, diverged about 268 ± 68 million years ago.  This would put the duplication event, under the tetraploidization hypothesis, at the time that mammals started to diverge.  This result is inconsistent with the occurrence of a second tetraploidization event prior to the evolution of teleost fish and the presence of aldolase C in fish.  This may indicate differences/changes in the molecular clock or a different origin for aldolase C in fish.  These studies are continuing in the Tolan laboratory.
Figure of the vertebrate tree.  The root is Choradata and the most primitive living vertebrate is amphioxius.

In addition to gaining a clearer understanding of when any one of the currently known aldolase isozymes arose, aldolase sequence data obtained from various primitive fish and skates can be used to help distinguish which is the closest living sister group of the vertebrates.  Furthermore, the analysis may discover a predicted fourth aldolase, aldolase D, its ancestry and a possible reason for its silencing in mammals.
Evidence for Tetraploidization
Gene duplication is thought to be important in the generation of the genetic diversity that determines the adaptability of a species to a changing environment [212,213].  Muller (1935) [213] and Haldane (1932) [212] first suggested that a redundant duplicate of a gene may acquire divergent mutations and may eventually emerge as a new gene.  Two explanations have been proposed for the existence of multiple copies of genes.  One explanation is that copy number increased by tandem duplication [211].  This could occur by; (1) unequal crossing over between homologous chromosomes in meiosis, (2) unequal exchange between sister chromatids during mitosis, or (3) regional redundant replication of DNA.  In all cases, tandem duplication leads to duplicate linked genes.  The second explanation is that gene duplications occurred by way of one or more tetraploidization events [207,209,210].  A heritable state of tetraploidy may arise when two diploid (as opposed to the normal haploid number) gametes fuse during fertilization and create a viable and fertile offspring.  The offspring, when mated to another tetraploid, which arose by a similar mechanism, would give rise to a new tetraploid lineage.
Actual examples of tetraploidization have been shown in some species of salmoniform fish.  Some salmoniform fish have a DNA content about 40% that of mammals, whereas other salmoniform fish such as the rainbow trout (Salmo irideus) have a DNA content about 80% that of mammals.  As a concrete example, two copies of the lactate dehydrogenase (LDH) gene are found in smelt (Hypomesus pretiosus) and the rainbow trout possess four copies [207].
In 1970, Susumu Ohno proposed that vertebrates arose as a result of two tetraploidization (genome duplication) events [209].  These events were speculated to have occurred (1) after that first primitive chordates arose (~500 Mya) and (2) about the time that the tetrapods (four-limbed vertebrates) arose (~375 Mya).  Several pieces of evidence support the tetraploidization theory.  Genome sizes (amount of DNA per haploid cell) of organisms from various vertebrate and chordate taxa revealed a pattern suggestive of two genome duplication events during vertebrate evolution: one prior to the evolution of the jawless fish and one before the evolution of the tetrapods [209].  Although this is generally true, DNA content is only a very rough measure of gene number.  In addition, however, the observation that the gene loci for the a- and b- chains of human hemoglobin were located on different chromosomes suggested that tetraploidization must have been the mechanism responsible for vertebrate evolution [207].  This hypothesis assumes that vertebrates evolved from a primitive chordate that had a relatively small number of gene loci.  However, the hemoglobin gene family is only one example consistent with tetraploidization.  There is solid evidence from a number of gene families this process as outlined below:
xMore evidence for the tetraploidization hypothesis came from the location of four human aldolase genes .  The aldolase genes are found on separate chromosomes.  More importantly, the chromosomes on which they were found are thought to be syntenous [214].  The relationship between the chromosomal locations of the human aldolase genes in the context of the tetraploidization theory can be seen in Figure x.  The branching in the trees corresponds to the tetraploidization events.  The current chromosomal locations and percentage differences in protein sequences are given for the human aldolases A, B, & C.  For the pseudogene on chromosome 10, the percentage difference is between it and aldolase B in the flanking regions .  The similarity in sequence in one pair of isozymes A & C, and the similarity in sequence of the gene flanking regions in aldB and the pseudogene locus suggests that two tetraploidization events is the most parsimonious explanation for the origins of these four aldolase genes.  Furthermore, the discovery of an aldolase C gene in goldfish [110], a teleost fish, suggests that a second tetraploidization event occurred, much earlier than first proposed [209].

Figure.  Evolutionary trees of aldolase genes.  Panel A.  The evolutionary tree illustrates the proposed evolution of the chromosomes containing aldolase loci predicted by chromosomal locations.  The arrows point to tetraploidization events as predicted by Ohno (1973) [208].  The scale is in millions of years.  Panel B.  A tree based on a difference matrix, including correction for multiple mutations at the same position (Fitch & Margoliash, 1967), was derived from the protein sequences.  The numbers are the percentage of relative difference along each limb.  Panel C.  A tree based upon alignments that can be made between flanking regions at the aldolase C, B, and pseudogene loci on chromosomes 17, 9, and 10, respectively.  The scale is in percentage difference.  The grey limb for aldolase A is presumed, as no data exists.  Taken from .
Lactate Dehydrogenase (LDH)
Quattro et al. (1993) showed that LdhA and LdhB evolved by duplication of a primordial Ldh locus after the origin of lamprey .  The two genes are found on different chromosomes in humans, 11 & 12 [215].  Interestingly, lamprey has been shown to have only one LDH, whereas all other vertebrates have at least two (the A and B isozymes).  Furthermore, human chromosomes 11 & 12 are thought to share considerable synteny [214].  Some vertebrates (actinopterygian fish, birds of the family Columbidae, and mammals) have independently gained a third isozyme, LDH C, by tandem duplication (e.g., in humans LDH A and LDH C are encoded on human chromosome 11 p15.4-p14.3 & 11 p15.1-p14.1, respectively [215]).
Hox Genes
Garcia-Fernandez and Holland (1994) found only one Hox gene cluster in the cephalochordate amphioxus [216].  Only one cluster has been found in all insects examined.  (Previous studies indicating two Hox gene clusters in amphioxus were questioned by Garcia-Fernandez and Holland.) Mice have four Hox-gene clusters [217] and lamprey have three clusters .  The above data are an indication that tetraploidization has occurred at least once during vertebrate evolution.
The sequencing of many genomes has revealed a process of shuffling of genes between chromosomes, which has clouded this kind of analysis.  The roles that tetraploidization versus tandem duplication versus shuffling versus gene loss have played in the evolution of the vertebrate genome is still under investigation and is somewhat controversial .
Previous studies have studied the evolution of a specific gene family without looking at the implications that this information has on the larger picture of genome evolution.  If tetraploidization was important in vertebrate evolution, the relationship of the descendent orthologous genes can be predicted provided they are subject to sufficient evolutionary constraints to have maintained sequences indicating their ancestry.  The aldolase gene family is suitable for such studies because, like the Hox cluster, there are more than two members in present vertebrates and members of this gene family are known to exist on separate syntenous chromosomes, at least in humans.

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.Phylogenetic analysis of the aldolase genes identifies isozyme specific residues
The formation of families of evolutionarily related but functionally distinct genes during vertebrate evolution, due to various mechanisms of gene duplication and subsequent divergence, is a fundamental process of adaptive evolution.  The mechanisms by which these processes occur are not completely understood.  To determine what particular mutations in a set of diverging genes gave rise to enzymes with similar function but different activities (i.e., isozymes) one can use phylogenetic analysis.  Understanding the roots of the often-subtle differences in the structure/function of isozymes is difficult, but lends itself to this analysis.  The relationship of functional differences to the less obvious structural differences among isozymes of a given gene family is difficult to elucidate by structural information alone.  We are interested in analyzing primitive chordate aldolases using phylogenetic techniques in order to identify amino acid residues in the aldolase isozymes that either; (a) are functionally important and lead to the distinct kinetic differences and physiological roles of the aldolases [110,111,202,225], or (b), have been fixed by random genetic drift, by virtue of being unselected or neutral [183,226].  The identification of isozyme-specific residues (ISRs) and the restricting of that group of conserved residues have given insights into their functional roles [111,225].
We have analyzed the vertebrate aldolase genes in many species of vertebrates and done both phylogenetic analysis and reconstruction of ancestral genes/proteins.  There are two avenues of inquiry into how the vertebrate genome has evolved.  The first is to continue to investigate the aldolase genes of lower vertebrates in primitive fish and other species and analyze these species for the number of aldolase genes and their phylogenetic relationships using computational approaches.  The second avenue is to use the sequence databases, look for isozymes from other vertebrate species, and investigate their relationship to one another.  For those for whom there are two or more distinct genes, the ancestral proteins will be generated computationally and from there the divergence times can be estimated.  If done on enough species, it can determine if many/all vertebrate isozyme genes diverged at different times or the same time.  If at the same time, it lends support for the tetraploidization hypothesis.  If at different times, it lends support for the tandem duplication and gene-shuffling hypothesis as a mechanism of molecular evolution.

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