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Does the Urea Cycle exist in invertebrates?

Does the Urea Cycle exist in invertebrates?



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Do invertebrates (like Drosophila and C.elegans) have a urea cycle?


C. elegans is missing all of the urea cycle enzymes. Drosophila is missing just one enzyme, but there's a shortcut which may enable it to nonetheless complete the cycle. (For comparison, here's the pathway map for human with the complete cycle.)


Perhaps worth adding that the role of the urea cycle in terrestrial vertebrates is the disposal of excess N derived from protein metabolism. Such animals are ureotelic. Arthropods, in contrast, are uricotelic, using uric acid for N disposal, so it would make no sense for them to have a functional urea cycle. (Nematodes - I don't know.)

The importance of avoiding having a urea cycle when it is not needed is illustrated by the situation in yeast (Saccharomyces cerevisiae). In yeast some of the enzymes that would be part of the urea cycle are used in degrading arginine, while others are used in the biosynthesis of arginine. Yeast cells will carry out one or the other of these processes depending upon their metabolic status. If both processes were to be active at the same time this would result in a urea cycle that would waste ATP. This is avoided by having a multienzyme complex consisting of arginase and ornithine transcarbamoylase (in the degradative and biosynthetic pathways respectively), allowing reciprocal regulation so that only one of these is active at any time.


Ammonium

The ammonium cation is a positively charged polyatomic ion with the chemical formula NH +
4 . It is formed by the protonation of ammonia (NH3). Ammonium is also a general name for positively charged or protonated substituted amines and quaternary ammonium cations ( NR +
4 ), where one or more hydrogen atoms are replaced by organic groups (indicated by R).

  • 14798-03-9 Y
  • 54S68520I4 Y

Background to the protocol

What is urea?

Every organism decomposes nucleic acids and proteins, generating nitrogenous waste because nucleic acids and proteins contain nitrogen. Mammals, amphibians and some invertebrates excrete nitrogenous waste as urea, which is produced in the liver. Urea is an especially good compound for disposing of nitrogen because it is water-soluble and less toxic than ammonia – the excretory produce of fish, for example. Human urine contains 2% urea.

Urea was also the first organic compound ever synthesised. In 1828, Friedrich Wöhler synthesised urea from inorganic compounds (lead cyanate and ammonium hydroxide). This was a landmark achievement: until then only living organisms were believed to be able to produce organic compounds, and these compounds were thought to be special and to require a ‘vital force’ to make them. Wöhler bridged the gap between the living and non-living worlds. He didn’t receive a Nobel Prize for his discovery though, because the Nobel Prize did not exist at that time. Today, urea is synthesised in vast quantities: it is used to make plastics and as a cheap nitrogenous fertiliser.

What is urease?

Urease catalyses the hydrolysis of urea to carbon dioxide and ammonia. It is found mainly in seeds, micro-organisms and invertebrates. In plants, urease is a hexamer – it consists of six identical chains – and is located in the cytoplasm. In bacteria, it consists of either two or three different subunits. For activation, urease needs to bind two nickel ions per subunit.

How did urease become famous?

Urease from jack beans (Canavalia ensiformis) was the first enzyme ever purified and crystallised, an achievement of James B. Sumner in 1926, at a time when most scientists believed that it was impossible to crystallise enzymes. This earned Sumner the 1946 Nobel Prize in Chemistry. Today, crystallisation of proteins helps scientists to discover their structure and determine how they work. This knowledge permits the design of substances that interfere with enzyme action, such as the anti-AIDS drugs which inhibit the action of HIV’s enzymes or recent developments towards a possible rabies treatment (Ainsworth, 2008).

Why is there urease in soya beans?


The soya plant
Image courtesy of iStockphoto

The role of urease in soya beans is not entirely clear, although it is possible to speculate. Soya leaves also contain urease, but here, the enzyme is a thousand times less active than in the beans. It is known that the leaf enzyme helps to recycle nitrogen from proteins (the proteins are broken down to urea). In the beans, urease does the same when the beans germinate. The resulting ammonia from the reaction may also protect the plant cells from pathogens – it seems that the plant enzyme itself is an insecticide.

Where else can urease be found?

Many species of bacteria produce urease, including Helicobacter pylori, the bacterium responsible for stomach ulcers. By doing this, H. pylori raises the pH of the gastric juice from about pH 3 to pH 7, the optimal pH for its growth. A commercially available test for H. pylori checks for the presence of urease in breath, and is used as a tool for diagnosing stomach ulcers.

Ruminants (such as cows and sheep) have cellulose-digesting bacteria in their rumen – the first compartment of their stomachs – to help them digest their plant diet. Ruminants excrete urea into this part of the stomach, the urea making an excellent source of nitrogen for bacterial growth. To take up the nitrogen, the bacteria secrete urease to break down the urea. Eventually, the animals digest the bacterial mass.

Is urease in soya beans harmful to humans?

Urease is not harmful. However, raw soya beans contain other compounds which are unhealthy. For example, there is a protein inhibitor in raw soya beans which prevents the digestive enzyme trypsin from working and makes raw soya beans inedible. The presence of the inhibitor is not easy to detect, but fortunately, it has a similar level of heat intolerance to urease – both are inactivated by heating. Therefore, to ensure that the inhibitor is inactivated, commercial preparations of soya beans (soya flour or foods that contain soya, such as tempe and tofu) are tested for urease activity – in a very similar way to the test described here. If no urease activity can be detected, the inhibitor has presumably also been inactivated.

Urease in the nitrogen cycle

Nitrogen is a crucial element for plant growth, but most plants can only use it in the form of ammonium or nitrate. Only legumes (thanks to the bacteria they live in symbiosis with) and cyanobacteria can use elemental nitrogen from the air.

Many animals excrete urea in their urine. Soil micro-organisms feed on animal urine, producing urease to transform the urea to ammonia, which is then readily accessible to plants. This is part of the nitrogen cycle, the process by which the nitrogen from proteins and other compounds is constantly recycled.


Elimination Products of Nitrogen Metabolism | Biochemistry

We have seen that some amino acids are utilized by the animal organism as detoxication agents and that products like benzoyl-glycine (hippuric acid) or phenyl-acetyl-glutamine can be found in urine. We will also see that urobilin and stercobilin, the degradation products of hemoglobin (see fig. 7-33), are present in faeces.

But generally, nitrogen is eliminated in 3 forms:

1. Ammonia and ammonium salts in ammoniotelic organisms (aquatic in­vertebrates, fishes)

2. Uric acid, in uricotelic species (terrestrial invertebrates, birds, reptiles)

3. Urea in ureotelic animals (especially mammals including man).

These various nitrogeneous wastes can co-exist in the same organism for example, man excretes nitrogen mostly in the form of urea (80%), but am­monium salts and uric acid are also found in urine.

Ammonia is a highly toxic product the organism must be able to eliminate it fast and that is probably why this form of elimination was maintained in aquatic animals where NH3 can be expelled rapidly by diffusion into the external medium.

In terrestrial animals less toxic products are used: uric acid and urea. This idea is confirmed by the fact that some animals (e.g., the tadpole) excrete their nitrogen in the form of NH3 during the aquatic period of their life, but acquire the property of forming urea when they leave water (frog).

1. Ammonia and Ammonium Salts:

As observed above, in most tissues, the deamination of amino acids leads to the formation of NH3. This toxic ammonia is taken over by glutamic acid to form glutamine and it is in this form that NH3 is transported by blood up to the kidneys.

The glutamic acid-glutamine system therefore plays a capital role in the transport of NH3, and it must be mentioned that these 2 amino acids represent a very high percentage of the non-protein nitrogen of animal tissues (in human plasma they together form 1/3 of the total nitrogen of amino acids).

In the kidneys, glutaminase catalyzes the reverse reaction and decomposes glutamine which thus provides the major part of urine NH3. The rest is produced by the deamination of amino acids in the renal parenchyma. In the liver, the ammonia produced by glutamate dehydrogenase is directly detoxified in the urea cycle by a mechanism we will study in the next paragraph.

In the case of acidosis, for example during prolonged physical exercise (production of lactic acid), or in diabetes (ketonemia), or after injection of acid to an animal, excretion of NH3 increases – at the cost of urea formation — because it is used for neutralizing acidity, which permits an economy of other ions required by the organism (Na + , Ca 2+ , Mg 2+ ) therefore, in acidosis, more NH3 is eliminated and less urea.

Exactly the reverse takes place in the case of alkalosis. It may be said that in general, in an animal receiving a fixed diet, the total nitrogen excreted ( NNH3 + Nurea) does not vary much, but the respective proportions of the two forms of elimination can vary.

As we will see now, NH3 is a precursor of urea, but it must remembered that the reverse is not true (urea is not a precursor of urine NH3, the precursor is mostly blood glutamine, and — to a lesser extent — all the amino acids whose deamination in the kidney produces NH3 which is immediately excreted).

2. Ureogenesis:

Urea formation is localized in the liver this was shown by various observa­tions and experiments and especially the fact that if an isolated liver is perfused with a solution containing NH3, the liquid flowing out is free from NH3 but contains urea. Krebs and Henseleit succeeded in bringing about urea forma­tion by slices of rat liver and showed that arginase catalyzes urea formation from arginine.

But there was not enough free arginine to explain urea produc­tion by the hydrolysis of this amino acid, and this led to the idea that arginine acts catalytically it was later found that two other amino acids, ornithine and citrulline have the same action and Krebs proposed the mechanism represented in figure 7-30, known as ornithine cycle, urea cycle, or Krebs-Henseleit cycle, the various steps of which we will examine now.

A. Synthesis of Carbamyl-Phosphate and Transformation of Ornithine into Citrulline:

The formation of carbamyl phosphate is catalyzed by a mitochondrial en­zyme using NH3 as substrate (the latter results from the deamination of glutamate by glutamate dehydrogenase).

A derivative of glutamate, acetyl- glutamate, is the allosteric activator of this carbamyl-phosphate synthetase (the carbamyl-phosphate required for the synthesis of pyrimidines is synthesized by a cytosolic enzyme, using glutamine as substrate and being regulated in a totally different manner).

Carbamyl-phosphate reacts with the δ-amino group of ornithine to give citrulline, under the influence of ornithine-carbamyl-transferase. It must be noted that during this step, car­bamyl phosphate supplies the carbon atom and one of the nitrogen atoms of the future urea molecule.

B. Transformation of Citrulline into Arginine:

This transformation requires two reactions. In a first step, citrulline con­denses with aspartic acid in presence of ATP to form arginosuccinic acid, which, in a second step is split into arginine and fumaric acid, it may be observed that the aspartic acid supplies the second nitrogen atom of the future urea molecule.

This transfer in two steps of the amino group of aspartic acid (which is converted into fumaric acid in the process) is quite similar to the transfers we studied, on one hand during the input of N, in the biosynthesis of the purine ring, and on the other hand during the amination of IMP to AMP.

The immediate donor of NH2 is therefore obligatorily aspartic acid, but we have seen that the amino group of aspartic acid may result from glutamic acid (by transamination to oxaloacetic acid) and that the amino group of glutamic acid may itself result from a large number of amino acids since glutamic acid play a very active part in transamination processes. The 2NH2 of the urea molecule may therefore originate from various amino acids.

C. Transformation of Arginine into Ornithine and Urea:

This hydrolysis is catalyzed by arginase and leads to urea on one hand and ornithine on the other ornithine is thus regenerated and can begin a new turn of the cycle by binding a new carbamyl-phosphate molecule.

D. Relations between the Cycle of Urea and the Cycle of Tricarboxylic Acids:

It is observed that the amino group of amino acids can be eliminated in the form of urea through 2 pathways which join in the ornithine cycle: either by deamination and then incorporation in carbamyl-phosphate, or by transamina­tion to aspartic acid (with a possible intermediate passage through glutamic acid).

In the latter case, it is apparent that there is a close relationship between the urea cycle and the cycle of tricarboxylic acids, thanks to aspartic acid and fumaric acid which establish the link between the 2 cycles: the oxaloacetic acid of the cycle of tricarboxylic acids can be transaminated, and the aspartic acid thus formed enters the urea cycle the fumaric acid thus formed returns to the cycle of tricarboxylic acids.

Furthermore, transamination which gives rise to aspartic acid often takes place at the cost of glutamic acid, which is itself formed from α-ketoglutaric acid, another intermediate of the cycle of tricar­boxylic acids (see fig. 7-31).

Besides, energetic relations exist between the 2 cycles since the synthesis of urea requires ATP for the synthesis of carbamyl-phosphate and also for the condensation of citrulline and aspartic acid, and the ATP required can be supplied by the phosphorylations which accompany the oxidation of the inter­mediates of the cycle of tricarboxylic acids. Krebs and Henselheit had in fact observed that the synthesis of urea in their experiments necessitated the presence of oxygen and the addition of an oxidizable substrate.

3. Uric Acid:

In uricotelic animals like birds, uric acid is the principal form of elimination of nitrogen it is a compound possessing the purine ring which is synthesized from various precursors. These animals therefore perform a series of relatively complex reactions to eliminate their nitrogen.

In man, uric acid is also found in urine it results from the catabolism of purine nucleotides (which are themselves produced by the hydrolysis of nucleic acids). Figure 7-32 shows the catabolism of the two purine bases, adenine and guanine, which appear after the action of nucleosidases on nucleosides (them­selves formed by action of nucleotidases on nucleotides).

It is observed that the two bases are first deaminated, then oxidized thanks to an enzyme called xanthine-oxidase which catalyzes the transformation of xanthine into uric acid and functions with a flavin coenzyme transferring the hydrogen atoms directly to the oxygen.

In man, the catabolism of the purine ring ends there, uric acid is eliminated in urine. In a disease called gout, uric acid salts (urates) of low solubility, are deposited in the joints.

But in most animals the catabolism of the purine ring continues (see fig. 7-32) up to the allantoin stage (in many mammals), the allantoic acid stage (in some fishes), or even up to the urea and glyoxylic acid stage. Lastly, let us mention that some bacteria possess an enzyme, urease, which can hydroiyze urea to CO2 and NH3.

4. Catabolism of Porphyrins:

The porphyrin group of hemoglobin remains stable for the entire life of the erythrocyte (about 4 months), as maybe noted by administering the 15 N isotope whose level in the prosthetic group remains constant during this period.

Then, hemoglobin is degraded mostly in the spleen — up to the bilirubin stage (see fig. 7-33) bilirubin is then transported to the liver where it is conjugated with glucuronic acid, before being eliminated through the bile, into the intestine.

When elimination of the bile is prevented by an obstacle (e.g., a calculus), the bile constituents flow back into the blood, particularly the glucorono-conjugated derivative of bilirubin, which can then pass into the urine. But when bile flows normally into the intestine, bilirubin reaches there and undergoes a new series of transformations under the influence of intestinal bacteria and their enzymes.

Free bilirubin is first regenerated by hydrolysis of the glucorono-conjugate and then the following may take place (see fig. 7-33):

1. Reduction of the 2 vinyl groups (on pyrrol I and pyrrol II) to ethyl groups, which gives mesobilirubin

2. Reduction of the 2 β and δ — CH2= bridges to — CH3 —, which gives mesobilirubinogen (or urobilinogen).

There are then 2 possibilities:

1. Either dehydrogenation of the γ methylene bridge and the nitrogen of pyrrol IV, which gives urobilin,

2. Or reduction of the double bonds of pyrrol I and pyroll II, which gives stercobilinogen which is transformed into stercobilin by a dehydrogenation of the γ methylene bridge and the nitrogen of pyrrol IV.

Urobilin and stercobilin are excreted in the faeces which take up their coloration (this is why stools are sometimes discoloured during disorders of bile secretion). As for uribilinogen and stercobilinogen, they are partly re-ab­sorbed by the intestinal mucosa and return to the liver an entero-hepatic cycle of bile pigments is thus produced.


Materials and Methods

Ethics Statement

Experimental procedures involving animals were approved by the Institutional Animal Care and Use Committee of the Children’s National Medical Center.

Determination of the Solvent Accessible Surface Area and Conservation

Crystal structures 5DOT, 5DOU, and 1OTH were used to calculate relative solvent accessible surface area (SASA) for the apo and liganded CPS1, and OTC trimer structures after removal of heteroatoms and water molecules. SASA of each amino acid was calculated with the Shrake and Rupley dot method (Shrake and Rupley, 1973) as described by Ho 1 and using mesh density 9,600. A custom Python script 2 was used to calculate SASA for each residue. The same method was used to calculate maximal SASA for amino acid using polypeptide in which each of the 20 amino acids is flanked by a glycine residue (Supplementary File S1) this polypeptide was modeled as β-strand using VEGA 3.1.1 (Pedretti et al., 2002). Relative SASA was calculated by dividing SASA of each amino acid with its maximal SASA.

Conservation of amino acids was determined from the alignment of either 233 homologs of human CPS1 (Supplementary File S2) or 270 homologs of human OTC (Supplementary File S3) from vertebrates and multicellular invertebrates. Protein sequences were collected from the NCBI non-redundant protein sequence database using Protein BLAST (Altschul et al., 1990, 1997), default parameters (word size 6, expected threshold 10, scoring matrix BLOSSUM62, gap existence 11, and gap extension 1) and sequences of human CPS1 and OTC as queries. Clustal Omega (Madeira et al., 2019) was used for multiple protein sequence alignment and WebLOGO3 (Crooks et al., 2004) was used for visualization of multiple sequence alignments. Conservation of surface residues that are mutated in patients with CPS1 and OTC deficiencies was determined as percent of either 233 CPS1 or 270 OTC homologs that have the same amino acid as human protein at that position.

Accurate Modeling of the Impact of Mutations

The structural models of CPS1 and OTC used in the prediction of the effects of point mutations were obtained after several modeling steps. First we modeled the missing loops, side-chains and termini into the existing structures of CPS1 and OTC (PDB entries 5DOU and 1C9Y, respectively) using MODELLER version 9.23 (Eswar et al., 2006). Arginine 270 in the crystal structure of OTC was reverted to glutamine according the sequence reported in the UniProtKB database (entry P0048) (UniProt Consortium, 2019).

The prediction of changes in protein stability (the ΔΔG) and structure resulting from single amino acid substitutions was performed with the ddg_monomer application, as implemented in Rosetta version 3.11, following the high-resolution protocol (Kellogg et al., 2011). The protocol generated 50 models for both the wild-type and the point mutant. The ΔΔG of the mutation was calculated as the difference in Rosetta energies between the three highest scoring wild−type structures and the three top−scoring mutant structures. Input structures were pre-minimized to reduce steric clashes. Distance restraints between Cα pairs within 9 Å of each other were part of the optimization to prevent the backbone from moving too far from the starting conformation. The ideal value for the restraint was taken as the distance in the original structure and the standard deviation on the harmonic constraint was set to 0.5 Å. The score12 weight function (Rohl et al., 2004) was used in all calculations. The crystallographic threefold symmetry was explicitly imposed on all OTC models both in MODELLER and Rosetta. Mutant proteins with ΔΔG of 0𠄲 kcal/mol were considered to have similar stability as the wild-type while mutant proteins with ΔΔG < 0 kcal/mol were considered to be more stable than the wild type protein.

Identification and Computational Analysis of VS Sequences

Protein sequences of vertebrate NAGS were collected using Blastp to query vertebrate proteins in either NCBI nr or UniProt databases with human and zebrafish NAGS (Caldovic et al., 2002a, 2014). Default parameters (word size 6, gap opening and extension penalties 11 and 1, respectively, and BLOSUM62 scoring matrix) were used for the search, which resulted in 90 mammalian NAGS sequences (Supplementary File S4) and 61 NAGS sequences from fish, amphibians and reptiles (Supplementary File S5). The most likely translation initiation site for each NAGS sequence was determined by inspection of protein alignments with the corresponding genomic sequences and with human and zebrafish NAGS, performed using ClustaW in MEGA7 (Kumar et al., 2018) amino acids encoded by predicted exons located upstream of the exon that corresponds to exon 1 in human and zebrafish NAGS genes were removed. The boundaries of the VS were defined as sequences between the mitochondrial protein peptidase (MPP) cleavage site and the beginning of sequence homology with vertebrate-like N-acetylglutamate synthase-kinase from Xanthomonas campestris (XcNAGS-K), which does not have MTS and VS (Qu et al., 2007). Sequence alignments with mouse NAGS, which has experimentally determined MPP cleavage site (Caldovic et al., 2010), as well as MitoPorotII (Claros and Vincens, 1996) and MitoFates (Fukasawa et al., 2015) were used for prediction of MPP cleavage sites in NAGS sequences. The C-termini of VS were determined by sequence alignments of NAGS sequences with XcNAGS-K using ClustalW in MEGA7. The lengths, proline content and sequence identities of VS were determined using MEGA7. WebLOGO3 (Crooks et al., 2004) was used to visualize VS sequence alignments that were generated with ClustalW in MEGA7.

Fractionation of Rat Liver Mitochondria

Fractionation of mitochondria was carried out as described previously (Powers-Lee et al., 1987). Briefly, mitochondria were purified from donated rat livers using differential centrifugation (Graham, 2001). Purified mitochondria were resuspended in 5 mM Tris HCl, 250 mM Sucrose, 1 mM EDTA, pH 7.2, and subjected either to three rounds of freezing and thawing, or treatment with 0.12 mg of digitonin per mg of mitochondrial protein to remove the outer mitochondrial membrane as supernatant after centrifugation at 9000 × g for 10 min. Pelleted material was resuspended in 20 mM Hepes Buffer, pH 8.0 and sonicated. The vesicles that resulted from the sonication treatment were treated with increasing concentrations of Triton X-100 (0, 0.1, 0.5, and 1.0%) for 30 min. at room temperature, followed by pelleting of the membranes by ultracentrifugation at 144,000 × g for 60 min, washing three times with 20 mM Hepes, pH 8.0, and re-suspension in the same buffer. The amount of NAGS in each mitochondrial fraction was determined using immunoblotting with the primary antibody raised against recombinant mouse NAGS at 1:5,000 dilution and HPRT-conjugated donkey anti-rabbit secondary antibody (Pierce) at 1:50,000 dilution. NAGS bands were visualized using SuperSignal West Pico kit (Pierce) according to the manufacturer’s instructions. The amounts of CPS1 and OTC in each mitochondrial fraction were determined using immunoblotting with primary antibodies raised against CPS1 or OTC at 1:5,000 dilution, followed by the HPRT-conjugated secondary antibody at 1:10,000 dilution. CPS1 and OTC were visualized using ECL Western Blotting Substrate (Pierce) according to the manufacturer’s instructions. The intensity of each band was measured using a GS-800 Calibrated Densitometer (Bio-Rad) and the Quantity One software package (Bio-Rad). Mitochondrial fractions were probed with antibodies raised against mitochondrial markers of the IMM, mitochondrial matrix and outer mitochondrial membrane: CoxIV (Abcam) at 1:5,000 dilution, Grp75 (Stressgen) at 1:1,000 dilution and VDAC (Pierce) at 1:1,000 dilution (Da Cruz et al., 2003 Rardin et al., 2008, 2009). Filters were then probed with the HPRT-conjugated secondary antibody (Bio-Rad). The CoxIV, Grp75, and VDAC bands were visualized using ECL Western Blotting Substrate (Pierce).

Cloning of Recombinant Mouse Variable Segments

Mouse variable segment (mVS) coding sequence was subcloned using pNS1 plasmid (Caldovic et al., 2002b) as a template and primers 5′-GGG ACA TAT GCT CAG CAC CGC CAG GGC TCA C-3′ and 5′-AGG TGG ATC CTT ATT ATT ACC AGT GGC GTG CTT CC-3′ which amplify the sequence between codons 49 and 117 of the mouse NAGS preprotein coding sequence. The amplification conditions were: initial denaturation at 95ଌ for 3 min., followed by 25 cycles of denaturation at 95ଌ for 30 s, annealing at 60ଌ for 30 s and extension at 72ଌ for 30 s, and final extension at 72ଌ for 5 min. using Pfu Turbo Hotstart DNA polymerase (Stratagene). This amplification product was cloned into pCR4Blunt-TOPO (Invitrogen) producing TOPOmVS. The correct coding sequence was confirmed by DNA sequencing. Plasmid TOPOmVS was cleaved with NdeI and BamHI sites and subcloned into pET15b to create pET15bmVS.

The amino acid sequence of the reversed variable segment (revVS) was generated by reversing the amino acid sequence of mVS. The amino acid sequence of shuffled variable segment (shVS) was generated by dividing the sequence of mVS in the middle, then inter-digitating the amino acid sequences of the two halves. The coding sequences of revVS and shVS, including three stop codons at their 3′ ends and NdeI and BamHI restriction sites at the 5′- and 3′-ends, were chemically synthesized as mini-genes and inserted into pIDTSMART-KAN plasmid (Integrated DNA Technologies) followed by subcloning into pET15b bacterial expression vector to create pET15brevVS and pET15bshVS plasmids.

Recombinant Protein Purification

Recombinant NAGS was purified as described previously (Caldovic et al., 2006 Haskins et al., 2008). Briefly, plasmid pET15bmNAGS-M (Caldovic et al., 2006) was used for overexpression of mouse NAGS-M in E. coli. Pelleted cells were resuspended in Buffer A (50 mM potassium phosphate, 500 mM KCl, 20% glycerol, 10 mM β-mercaptoethanol, 0.006%Triton X-100, 1% acetone, pH 7.5) containing 10 mM imidazole and lysed with 40 mM n-octyl-β-d-glucopyranoside. Cell lysate was loaded onto Ni-NTA agarose column and recombinant NAGS-M was eluted with Buffer A containing 250 mM imidazole.

Recombinant mVS, revVS, and shVS were purified from cultures of transformed Escherichia coli C41(DE3) cells that were induced with the Overnight Express Autoinduction Kit System 1 (Novagen). Cells were pelleted and resuspended in Buffer A containing 10 mM imidazole. Lysozyme and phenylmethylsulfonyl fluoride were added to the final concentrations of 1 mg/ml and 0.1 mM, respectively. The cells were lysed with 40 mM n-octyl-β-D-glucopyranoside. DNAseI and RNAseA (0.1𠄰.5 mg/ml lysate) in 5 mM MgCl2 were added to remove nucleic acids by incubation at room temperature for 30 min. Cell lysate was cleared by centrifugation at 25,000 × g for 30 min at 4ଌ. A nickel-affinity column (GE Healthcare) was equilibrated with buffer A containing 10 mM imidazole. Cleared lysate was loaded onto the column at a flow rate of 0.3 ml/min. The column was washed with Buffer A containing 50, 125, 250, and 500 mM imidazole. The variable segments eluted between 250 and 500 mM imidazole. The protein size and purity were verified by Comassie blue staining following SDS-PAGE on the 16.5% Tris-Tricine Gel (Bio-Rad).

Mass Spectrometry Peptide Sequencing of Mouse Variable Segments

To confirm the identity of the purified mouse variable segments, they were excised from the 16.5% Tris-Tricine Gel and subjected to rapid, in-gel trypsin digestion (Shevchenko et al., 2006). The fragments were analyzed using mass spectrometry on an Applied Biosystems Voyager 4700 MALDI TOF/TOF mass spectrometer (Supplementary File S6).

Co-immunoprecipitation

Mouse liver mitochondria were purified from donated tissue using differential centrifugation (Graham, 2001) and lysed with PBS containing 2% CHAPS (Stankiewicz et al., 2005). Mitochondrial lysate was diluted to 1 mg/ml protein for immunoprecipitation with antibodies against OTC and CPS1 and 5 mg/ml protein for immunoprecipitation with anti-NAGS antibodies. Mitochondrial lysates were mixed with magnetic beads (Invitrogen) cross-linked to primary antibodies against NAGS, CPS1 or OTC according to the manufacturer’s instructions. Following incubation at 20ଌ for 10 min, the beads were washed five times with PBS containing 0.05% Triton X-100. Protein complexes were eluted with ImmunoPure IgG Elution Buffer (Pierce). Protein concentration in each elution fraction was measured using protein assay dye reagent concentrate (Bio-Rad) according to the manufacturer’s instructions. Between 0.5 and 1 μg of immunoprecipitated proteins were resolved using SDS-PAGE, and probed with primary antibodies raised against NAGS, CPS1, or OTC followed by the HPRT-conjugated secondary antibody. NAGS was visualized using SuperSignal West Pico kit (Pierce), and CPS1 and OTC were visualized using ECL Western Blotting Substrate (Pierce).

In experiments measuring competition between NAGS-M and the recombinant mVS, mitochondrial lysate was diluted to a protein concentration of 2 mg/ml, mixed with the mVS, revVS, or shVS in a 1:1 (v/v) ratio, and added to magnetic beads (Invitrogen) cross-linked to primary antibodies against CPS1. Depending on the experiment, the molar excess of recombinant variable segment peptides relative to CPS1 was between 10 and 30-fold, based on estimates of the reported abundance of CPS1 in the liver mitochondria (Raijman, 1976 Cohen et al., 1982 Sonoda and Tatibana, 1983 Wang D. et al., 2019). Immunoprecipitation was carried out as described above. The intensities of NAGS-M bands were measured using a GS-800 Calibrated Densitometer and Quantity One software (Bio-Rad).

Confocal and gSTED Microscopy

Confocal and Gated Stimulated Emission Depletion (gSTED) microscopy were performed as described previously (Bhuvanendran et al., 2014 Salka et al., 2017). Imaging was performed using the Leica TCS SP8 microscope equipped with a white light laser, two depletion lasers, acousto-optical beam splitter (AOBS) and hybrid detectors. Single labeling of all the confocal and gSTED samples was done using Alexa Fluor 647 while the double-labeled samples also had Alexa Fluor 532.

An HC PL APO CS2 100x/1.40 Oil objective was used to acquire confocal images. Alexa Fluor 532 was excited using 515 nm laser line and the emission was collected on a hybrid detector with the AOBS set to 520� nm whereas the Alexa Fluor 647 was excited using 645 nm laser line and the emission between 650 and 720 nm was collected.

12-bit gSTED images with pixel size less than 30 nm were acquired using the HC PL APO CS2 100x/1.40 Oil objective. Samples with Alexa Fluor 647 fluorophores were excited at 645 nm and depleted with 775 nm laser. The emission was collected between 650 and 720 nm with a time gating of 0.3𠄶.0 ns. In the double-labeled samples, sequential stack for Alexa Fluor 532 was acquired using a 515 nm excitation and 660 nm depletion. The time-gated emission between 2.2 and 6.0 ns was collected with the AOBS set from 520 to 590 nm.

These confocal and STED images were deconvolved with Huygens Professional version 17.04 (Scientific Volume Imaging 3 ). Further image analysis, including intensity plots, were done using MetaMorph Premier version 7.7.0 (Molecular Devices 4 ).


Effector Cells

The basic phagocytic ability of unicellular organisms (e.g., amebae) is also found in the most primitive multicellular animals belonging to the group Porifera (sponges) and cnidarians (the group including jellyfish and sea-anemones). These animals apply phagocytic amebocytes for nutrition and recognition of foreign elements in the environment. Similar cell types have been conserved through evolution as they are recognized in all groups from invertebrates (annelids, arthropods, mollusks, echinoderms) to vertebrates (4). Several terms have been assigned to these cells in various groups and it must be expected that future investigations will sub-divide groups further. Sponges carry amebocytes in their mesoglea, cnidarians possess interstitial cells with a phagocytic function, hemocytes are found in the vascular system, and coelomocytes occur in coelomate animals. Thus, earthworms possess several subtypes of coelomocytes including eleocytes, and granular amebocytes (5) and in arthropods, comprising both crustaceans and insects, several effector cell types have been characterized (19). The evolutionary importance of corresponding phagocytes/macrophages is reflected in the range of subsets described from invertebrates and primitive chordates. Various cell types within this theme are found in advanced invertebrates (represented by echinoderms such as sea stars and sea urchins) and in the cephalochordate Branchiostoma (Amphioxus) and in urochordates (tunicates, ascidians) where both granulocyte-like cells and macrophages occur (20, 21). An even more diverse array of cell types and subsets occur in jawless vertebrates (hagfish and lampreys), cartilaginous fish (sharks and rays), and in bony fish. Besides phagocytes, jawless fish possess different subsets of lymphocytes with special membrane receptors. These primitive vertebrates without jaws have evolved an alternative antigen recognition system, which are composed of LRRs. These molecules provide agnathans a basis for establishing various lymphocyte lines corresponding to B and T lymphocytes. However, in cartilaginous and bony fish, the lymphocyte receptors are immunoglobulin (B-cell receptors) or T-cell receptors whereas agnathans apply at least two forms of variable lymphocyte receptor (VLR) based on LRR (13).

In bony fish, the cellular armament might include lymphocytes, macrophages, monocytes, dendritic cells, neutrophils, granulocytes, eosinophils, basophils, mast cells, and NK-cells and an even higher specialization is known in mammals (6, 7, 22). Leukocytes have traditionally been divided into the myeloid and lymphoid line based on their development from certain stem cells. However, B-lymphocytes in rainbow trout have been shown to exert phagocytosis (23), which suggests that the border between these developmental cell lines is less rigid at least in fish. In this context, it is interesting that the Ikaros multigene family, which take part in vertebrate hemopoietic stem cell differentiation and production of B, T and NK cell lineages, seems to find an early version in the most primitive vertebrates (the agnathan hagfish Myxine) and the even earlier urochordates (the tunicate Oikopleura) (24). The ancient origin of genes, which are central in cellular adaptive immunity in higher vertebrates, is also reflected by the finding of a Nuclear Factor of Activated T-cells (NFAT)-like gene in the primitive chordate Branchiostoma belcheri (Amphioxus group). In this chordate, this gene seems to play a role in innate recognition of lipopolysaccharide (LPS).


Urea Production and Metabolism

Clinical Uses

Uremic symptoms are principally due to the accumulation of ions and toxic compounds in body fluids (79). Because protein-rich foods are the major source of these waste products, CKD can be considered a condition of protein intolerance. Indeed, it has been known since at least 1869 that restricting the amount of protein in the diet of patients with kidney diseases improves their uremic symptoms (80). More recently, we learned that dialysis efficacy is reflected in the removal of urea because changes in urea accumulation reflect changes in accumulated metabolic waste products. Again, this is not a new concept: The link between dietary protein and urea has been recognized since at least 1905, when Folin reported that urea excretion varies directly with different levels of dietary protein (81). These relationships were elegantly documented by Cottini et al. ( Figure 12 ), who fed patients with CKD different amounts of protein (expressed on the abscissa as nitrogen intake because 16% of protein is nitrogen) (82). With low levels of dietary protein (e.g., approximately 12 g protein/d equivalent to approximately 2.5 g nitrogen), nitrogen balance was negative, indicating that this level of dietary protein causes progressive loss of protein stores. When the diet was raised Ϥ g nitrogen/d, nitrogen balance became positive, signifying that protein stores were being maintained. With progressively more dietary protein, nitrogen balance remained positive but changed minimally. Instead, when dietary protein was above the level required to maintain nitrogen balance and protein stores, it was used to make urea. Clearly, urea production reflects the level of protein in the diet and the risk of developing complications of uremia. In addition, a high-protein diet invariably contains excesses of salt, potassium, phosphates, and so forth (83). The clinical problems that arise from high-protein diets in patients with CKD were recently highlighted in reports concluding that increases in salt intake or serum phosphorus will block the beneficial influence of angiotensin-converting enzyme inhibitors to delay the progression of CKD (84,85).

Urea excretion in adult humans with varying degrees of kidney malfunction fed milk, egg, or an amino acid mixture: assessment of nitrogen balance. Modified from reference 82, with permission.

Urea has special properties that can be used to evaluate the severity of uremia or the degree of compliance with prescribed changes in the diet. These properties include the following: (1) a very large capacity for hepatic urea production from amino acids, (2) urea is the major circulating pool of nitrogen and it crosses cell membranes readily so there is no gradient from intracellular to extracellular fluid under steady-state conditions, and (3) the volume of distribution of urea is the same as water (the urea space is estimated as 60% of body weight) (86�).

One clinically useful calculation is the steady-state SUN (SSUN), which reflects the severity of uremia because it estimates the degree of accumulation of protein-derived waste products. The SSUN calculation is useful because uremic symptoms are unusual when SSUN is 㱰 mg/dl.

The requirements for the calculation are that the patient with CKD is in the steady state (i.e., his or her SUN and weight are stable) and urea clearance in liters per day is known. Using the equation below, the amount of dietary protein that will yield a SSUN of 70 mg/dl can be calculated as:

The following steps are used to calculate the SSUN. First, the prescribed dietary protein in grams per day is converted into dietary nitrogen by multiplying the grams per day of dietary protein by 16%. Second, the nonurea nitrogen in grams of nitrogen excreted per day is calculated as the excretion of all forms of nitrogen except urea. This amount is approximated as 0.031 g nitrogen/kg per day multiplied by the nonedematous, ideal body weight (4,89). Third, the nonurea nitrogen is subtracted from the nitrogen intake to obtain the amount of urea nitrogen that must be excreted each day in the steady state. Finally, dividing the urea nitrogen excretion in grams per day by the urea clearance in liters per day yields the SSUN in grams per liter.

For example, consider a 70-kg adult with a urea clearance of 14.4 L/d (or 10 ml/min) who is eating 76 g protein/d. His SSUN (in grams per liter) is calculated from the following: 12.2 g/d dietary nitrogen−(0.031 g nitrogen/kg per day times 70 kg). The result is divided by the urea clearance in liters per day and multiplied by 100 to convert SSUN 0.69 g/L to 69 mg/dl.

This calculation arises from the demonstration that in the steady state, the production of urea is directly proportional to the daily protein intake ( Figure 12 ). The only other assumption is that urea clearance is independent of the plasma urea concentration, which is reasonable for patients with CKD. The key concept is that steady-state concentrations of nitrogen-containing waste product produced during protein catabolism will increase in parallel to an increase in the SSUN (4,82,89). By varying the amount of dietary protein, changes in the diet can be integrated with different values of the SSUN. As shown in Table 1 , similar concepts can be used to determine whether a patient is complying with the prescribed protein content of the diet (81,86,88).

Table 1.

Estimation of protein intake from urea metabolism

A 60-year-old man with stage 5 CKD is admitted to the hospital for plastic surgery. He weighs 70 kg and has been taught to follow a diet containing 40 g protein/d (6.4 g nitrogen/d because protein is 16% nitrogen). He excretes 4 g urea nitrogen/d, but on day 2 his BUN rises from 50 to 60 mg/dl.
 • The increase in BUN signifies accumulation of urea nitrogen in body water (70 kg x 0.6 L/kg x 0.1 g urea nitrogen/L = 4.2 g urea nitrogen/d).
 • His NUN is 70 kg x 0.031 g nitrogen/kg per day = 2.17 g nitrogen/d.
 • The total nitrogen excreted and accumulated is approximately 10 g/d (4 g urea nitrogen excreted/d +2.17 g NUN/d + 4.2 g urea nitrogen accumulated/d = 10.3 g nitrogen/d).
 • Because his nitrogen excretion substantially exceeds the dietary nitrogen of 6.4 g/d, he requires a consultation with a nutrition/dietician and testing for gastrointestinal bleeding

SUN, serum urea nitrogen NUN, nonurea nitrogen excretion.

These examples emphasize that the net production of urea in patients with CKD (also known as the urea appearance rate) can be used to estimate protein intake (4,82,89). For dialysis patients, the same relationships have been labeled as “urea generation” or the “normalized protein catabolic rate” (nPCR). Obviously, the nPCR equals the net urea production rate or the urea appearance rate except that it is not expressed per kilogram of body weight. However, the designation nPCR is misleading because the rates of protein synthesis and �tabolism” are far greater than the protein catabolic rate: The nitrogen flux in protein synthesis and degradation amounts to 45� g nitrogen/d, equivalent to 280� g protein/d (1). The principle of conservation of mass, however, indicates the difference between whole-body protein synthesis and degradation does estimate waste nitrogen production.

Urea Nitrogen Reutilization

Discussion of urea metabolism would be incomplete without addressing urea degradation. It is calculated from the plasma disappearance of injected [ 14 C]urea or [ 15 N]urea (88,90) and averages about 3.6 g nitrogen/d in both normal individuals and patients with uremia. The 3.6 g nitrogen/d arises from degradation of urea by ureases of gastrointestinal bacteria thereby supplying ammonia directly to the liver (88). Because this source of nitrogen could be used to synthesize amino acids and ultimately protein, the degradation of urea has been intensively studied (91). The evidence negates the hypothesis that urea degradation is nutritionally important. First, the amount of urea degraded has been expressed as an extrarenal urea clearance by dividing the rate of urea degradation by the SSUN. In normal adults, the extrarenal urea clearance averages approximately 24 L/d if this value were present in patients with CKD and a high SUN, the amount of ammonia derived from urea would be very high (79,88). However, the quantity of ammonia arising from urea in patients with CKD is not significantly different from that of normal individuals, indicating that the extrarenal clearance of urea in patients with CKD must be greatly reduced the mechanism for this observation is unknown (88).

Results from other testing strategies lead to the conclusion that it is unlikely that urea degradation contributes a nutritionally important source of amino acids to synthesize protein. We fed patients with CKD a protein-restricted diet and measured the turnover of urea using [ 14 C]urea. The results were compared with those obtained in a second experiment in which patients received neomycin/kanamycin as nonabsorbable antibiotics in order to inhibit bacteria that were degrading urea. In roughly half of the patients, antibiotic administration blocked urea degradation but there was no associated increase in urea appearance. This result means that ammonia arising from degradation is simply recycled into urea production and hence does not change urea appearance (90). We also addressed the hypothesis that removal of nitrogen released by urea degradation would suppress synthesis of amino acids and thereby worsen Bn. In this case, the hypothesis was rejected because inhibiting urea degradation with nonabsorbable antibiotics actually improved Bn (92). Finally, Varcoe et al. measured the turnover of urea and albumin simultaneously and concluded that the contribution of urea degradation to albumin synthesis was minimal (93).

The possibility that ammonia from urea degradation is used to synthesize amino acids was recently examined in hibernating bears (94). The authors noted that hibernating bears have very low values of SSUN (approximately 5� mg/dl) despite a decrease in GFR and they suggested that SSUN was low because urea was being used to synthesize amino acids. This finding would contribute to another oddity of hibernating bears, namely that their muscle mass and other stores of protein are relatively “spared” from degradation. Why the metabolism of hibernating bears might differ from that of patients with CKD is unknown and we applaud the investigators who gathered the information as experimenting on bears is quite tricky, even if they are hibernating.

Is Urea Toxic?

Because excess dietary protein produces uremic symptoms and because urea is the major source of circulating nitrogen, the potential for toxic responses to urea have been investigated using different experimental designs. Johnson et al. added urea to the dialysate of hemodialysis patients who were otherwise well dialyzed (95). Complications induced by the added urea were minimal until the SSUN was chronically 𾅐� mg/dl. This led to gastrointestinal irritation. There was no investigation of ammonia or inhibitors of urea degradation so the effect of urea can only be considered an association. An indirect evaluation of both mice with CKD and cultured cells revealed that urea may stimulate the production of reactive oxygen species. Reddy et al. concluded that a high SUN not only increased reactive oxygen species but also caused insulin resistance (96). However, it is difficult to assign insulin resistance to a single factor considering that there are so many uremia-induced complex metabolic pathways (97,98). It will be interesting to evaluate whether the production of reactive oxygen species initiates similar events in patients with CKD.

Another potential role of urea in producing uremia-induced toxicity is through the development of protein carbamylation, which could disrupt the structure of a protein interfering with signaling pathways and so forth. Stim et al. reported that the rate of carbamylation of hemoglobin increased in parallel with the increase in SUN and that carbamylation was significantly higher in patients with ESRD compared with normal individuals (99). These responses were confirmed by Berg et al. (100) except that the carbamylated protein was albumin, rather than hemoglobin. Thus, carbamylation of several proteins can occur in uremic individuals but whether this produces toxic reactions has not been defined.

Finally, there are patient-based reports that cast doubt on the hypothesis that urea is a toxin. Hsu et al. studied a man and a woman from a family of patients who had chronic but unexplained azotemia. Results of the evaluation indicated that the high SUN arose from a autosomal dominant genetic defect in urea reabsorption (101). Kidney function of the two participants revealed subnormal urea clearances but otherwise normal values of inulin clearance, urea excretion, and responses of urea clearance to diuresis and antidiuresis plus normal sodium clearances. Although the mechanism for the familial azotemia was not identified, the report is relevant because the participants had no clinical or laboratory findings attributable to the increase in SUN despite years of values varying from 49 to 65 mg/dl and from 55 to 60 mg/dl, respectively. In another case study, Richards and Brown studied a woman with prolonged azotemia to examine the association between a high SUN and the development of uremic symptoms (102). The participant subsisted on a diet consisting primarily of fish and a protein powder, yielding urea nitrogen production rates of 40� g/d for years. Although the participant maintained a SUN of 50� mg/dl for years, she had normal values of hemoglobin, plasma creatinine, BP, and no weight loss. Together, these reports indicate that even a prolonged increase in the concentration of urea does not produce toxic reactions, at least in patients with normal kidney function.

Urea is the largest circulating pool of nitrogen and its production changes in parallel to the degradation of dietary and endogenous proteins. These facts and other properties of urea can be used to estimate the degree of uremia and the compliance with prescribed amounts protein in the diet. The available evidence in patients with CKD suggests that reutilization of ammonia derived from urea degradation for the synthesis of amino acids and proteins is minimal. Whether the evidence would be more persuasive under extreme conditions, such as in hibernating bears, is unknown. The ability of urea to create toxicity is unsettled but years of high SUN values do not produce toxic reactions in individuals with otherwise normal kidney function.

In conclusion, renal urea and ammonia metabolism mediate critical roles in nitrogen balance, urine concentration, and acid-base homeostasis. In this review, we evaluated critical processes involved in these homeostatic mechanisms. Abnormal urea and ammonia metabolism both result from and can lead to a wide variety of conditions, including methods for evaluating issues that are critical to caring for patients with impaired renal function.


Conclusion and perspectives

The physiological considerations for the role of the UT family in vertebrates have been primarily concerned with the role of urea in concentrating urine in mammals, as well as in whole-animal urea excretion mechanisms in other lineages. However, the availability of molecular data both from a genomics point of view and from an expression standpoint provides us with new datasets that allow us to explore the evolution and function of UTs in vertebrate lineages.

Virtually all living cells have the metabolic machinery to be ureogenic. While a lot of attention has been focused on ureotelic vertebrates and urea production through the OUC, other ureogenic pathways are clearly important for homeostasis at the cellular and tissue level. For example, tissues such as the brain, testes or embryo proper are likely to rely on argininolysis to produce essential components for their function and proliferation, and in all these tissues, there is at least one UT mRNA expressed in most vertebrate species investigated to date. We therefore suggest that besides the association of the UT with typical ureogenic tissues (i.e. the liver via the OUC) and osmoregulatory and excretory tissues, it may be important to consider other ureogenic pathways (i.e. argininolysis) that are essential to tissue function to further understand the physiological role and evolutionary history of the UT in vertebrates. The need to remove urea from these tissues could be due to at least two potential pressures at the molecular level: (1) if urea is allowed to accumulate to levels high enough to lead to protein destabilization, an accumulation of a counteracting stabilizing co-solute (e.g. TMAO) is required (2) although the arginase reaction equilibrium is especially poised in the direction of urea production, accumulation of urea, a terminal end-product, cannot be allowed to occur to a level that might ‘back up’ these critical synthetic reactions via simple mass-action effects.

In addition to these physiological considerations, a number of important structural and functional questions remain. First, as mentioned earlier, there is little understanding of the multimeric structure of the UT in vertebrates. Although the structure of the UT-A1 (two UT single units in tandem) would suggest that the transporter could function as a dimer, the trimeric and dimeric structures of bacterial homologues could imply that different configurations are possible (Levin et al., 2009 Raunser et al., 2009). Further, the possibility of heterologous multimerization between the different UT isoforms has not been investigated to date. Although some tissues exhibit highly differentiated expression patterns, it is interesting to note that in a variety of species different UTs are co-expressed in a number of tissues. This co-expression could suggest that heterologous multimers of the transporter could indeed be functional and affect the properties and functionality of the multimeric UT.

The above brings us to the second elephant in the room: different UT isoforms may have different kinetics and properties that could generate transport capacities specific to the tissues in which they are expressed. As it is beyond the scope of the current review, we direct the reader to recent work discussing in more detail UT kinetics (MacIver et al., 2008 Stewart, 2011). In brief, it is established that the UT-A and UT-B proteins slightly differ in their susceptibility to phloretin, UT-B being more sensitive to this inhibitor (Martial et al., 1996). It is also suggested that UT-B has asymmetric transport capacities, promoting faster efflux of urea from red blood cells than the capacity for intracellular import (Macey and Yousef, 1988). These differences highlight a very important concept in UT characteristics, not only in terms of the overall capacity of these transporters but also in their respective roles and potential physiological function put forward previously. Indeed, in the case where a tissue or cell is better off iso-ureic with the external environment, then a UT-A-type transporter is probably a better choice to allow rapid equilibration of the tissue with plasma content. In contrast, a tissue that is more prone to damage by urea accumulation (e.g. testes, embryonic body) would need to quickly excrete urea rather than equilibrating to extracellular urea concentration and may use an asymmetric type of UT. The reasoning would be similar for tissues that reabsorb urea, for example the rainbow trout kidney, where an asymmetric transporter that facilitates the uptake of urea would allow a reduction in the urinary excretion of urea. In addition, some UTs may be more prone to localize to either the basolateral or apical membranes of epithelial cells, thereby conferring distinct permeabilities to these membranes and allowing urea flux in one or the other direction. While some advances have been made via immunohistochemistry to locate the UT in mammalian kidney (Bagnasco et al., 2001 Nielsen et al., 1996) and the fish gill (Bucking et al., 2013a,b), the cellular orientation of UTs on epithelial cells will continue to be a productive area of research for years to come.

Thus, now that we have examined the large UT repertoire in vertebrates, one can only wonder at the multiple possibilities due to not only the gene registry but also the multiple splice variants and potential for multimerization of these transporters and how these could affect urea handling in these animals. Hopefully, with modern techniques and the use of emerging model organisms, some of these questions can start to be addressed. At this point in time, our understanding of the UT physiological role across vertebrate taxa (excluding mammals) is only in its infancy despite the recent advances and productive efforts made in this field of research.


Urea-aromatic interactions in biology

Noncovalent interactions are key determinants in both chemical and biological processes. Among such processes, the hydrophobic interactions play an eminent role in folding of proteins, nucleic acids, formation of membranes, protein-ligand recognition, etc.. Though this interaction is mediated through the aqueous solvent, the stability of the above biomolecules can be highly sensitive to any small external perturbations, such as temperature, pressure, pH, or even cosolvent additives, like, urea—a highly soluble small organic molecule utilized by various living organisms to regulate osmotic pressure. A plethora of detailed studies exist covering both experimental and theoretical regimes, to understand how urea modulates the stability of biological macromolecules. While experimentalists have been primarily focusing on the thermodynamic and kinetic aspects, theoretical modeling predominantly involves mechanistic information at the molecular level, calculating atomistic details applying the force field approach to the high level electronic details using the quantum mechanical methods. The review focuses mainly on examples with biological relevance, such as (1) urea-assisted protein unfolding, (2) urea-assisted RNA unfolding, (3) urea lesion interaction within damaged DNA, (4) urea conduction through membrane proteins, and (5) protein-ligand interactions those explicitly address the vitality of hydrophobic interactions involving exclusively the urea-aromatic moiety.

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Section Summary

The breakdown and synthesis of carbohydrates, proteins, and lipids connect with the pathways of glucose catabolism. The simple sugars are galactose, fructose, glycogen, and pentose. These are catabolized during glycolysis. The amino acids from proteins connect with glucose catabolism through pyruvate, acetyl CoA, and components of the citric acid cycle. Cholesterol synthesis starts with acetyl groups, and the components of triglycerides come from glycerol-3-phosphate from glycolysis and acetyl groups produced in the mitochondria from pyruvate.


Watch the video: Urea cycle (August 2022).