Evolution

on August 22, 2009 in evolution

Today my friends and I went to Coronado to find the best evidence people have for evolution. No one could point us to a specific proof. We even talked to a scientist who talked about bacteria but offered no proof that one species transformed into another species? What is the true evidence that drives university teaching and television promotion of evolution?

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2 Responses to “Evolution”

  1. Jon says:

    [I wrote some of the following just now, but wrote the majority within the past two years.]

    The first step is in verifying the model of common ancestry. This can be done with some of the evidence given below. The next step is in determining the processes of biological change that account for it. It has long since been shown that natural selection acting on variation and the DNA that codes for it is the primary means of biological change. I’ll explain:

    Mutation adds variation to organisms’ preexisting genotypic variation. Configurations that increase fitness make the organisms more able (and thus more likely) to make significant contributions to their populations’ gene pools. Greater contributions increase the frequencies of the alleles with those configurations, with time. Likewise, the frequencies of alleles with configurations that decrease fitness decrease. That process is called natural selection, which acts on both preexisting and new mutation-derived variation. When a split breeding subpopulation (a deme) shifts into a new niche and/or environment, different configurations are then beneficial, detrimental, or neutral, so they genetically change in different ways. Since there is little to no gene flow between them and the greater population (either due to geographic location, changes in mating ritual, etc.), their changes are not shared. As the genetic difference increases, they can eventually no longer produce fertile offspring—and latter still, cannot produce any offspring.

    And there are many instances where this progression is quite apperent. For instance, there are island rings, where species on one island has radiated out, and each isolated subpopulation on each island can interbreed with the next, but one you come all the way around to the original island, the oldest deme cannot interbreed with the youngest.

    The same thing is observed in a salamander spices; as you look at demes from the mountain top, to the ones on either face, to the ones on either side of the base, each can breed with the one closest to it, but the ones either side of the base cannot interbreed.

    Even every singe creationist organization, such as Discovery Institute and Answers in Genesis, unambiguously state that natural selection is observable.

    In fact, natural selection is even observed to create protein-coding genes in labs. When the D2 domain of coliphage fd’s minor coat protein g3p (crucial for infectivity) was replaced with a random sequence of 139 amino acids and subjected to random mutagenesis. A 240-fold increase in fitness was observed after only 7 generations, eventually reaching a maximum of a 17,000-fold increase (Hayashi et al., 2006).

    But what about beneficial mutations? Are those also observed in labs? Absolutely. One of many observed examples of beneficial mutation is in the gene encoding the alpha isoform of the human tripartite motif-containing 5 antiviral protein (TRIM5?), as reported in the Kaiser (2007) study:

    TRIM5? is part of the intrinsic immune system, and restricts retroviral infection by binding with the retroviral capsid (Sebastian & Luban, 2005) and interfering with its uncoating process (Perron et. al., 2006).

    TRIM5 is comprised of a RING domain, a B-box domain, and a coiled-coil domain. The alpha isoform also contains a characteristic carboxyl-terminal SPRY domain (Reymond et. al., 2001)—also called the B30.2 domain. All of these domains play a roll in restriction (Javanbakht, et. al., 2005; Yap, Nisole, & Stoye, 2005); as does each of the three patches within the SPRY domain (Ohkura et. al., 2006). But the primary determinant of restriction specificity and efficiency is the first of the three patches (Sawyer et. al., 2004; Stremlau et. al., 2004).

    Armed with this information, in 2007, Dr. Shari Kaiser and her colleges set out to shed light on the distribution of type-1 Pan troglodytes endogenous retroviruses (PtERV1) among hominids by assessing the restriction efficiency of various primate TRIM5?s. The first road block they encountered was that “all copies of PtERV1 in the chimpanzee genome have been inactivated by accumulated detrimental mutations.” To circumvent this, they sequenced and aligned the capsid (CA) and p12 regions of various Catarrhine PtERV1 insertions, and constructed a consensus sequences to determine the original configuration. They then changed one these sequences to create a DNA copy of the original, and spliced it into murine leukemia virus (MLV), creating a PtERV/MLV chimeric virus.

    Viral vectors were used to insert TRIM5? genes into CrFK cells, resulting in their expression. They then infected those cells with the chimeras and with HIV1, and measured their efficiency of the TRIM5?s at restricting infection.

    They found that human TRIM5? is relatively inefficient at restricting HIV1, yet extremely efficient at restricting PtERV1.

    The unexpected part of the research came when they changed the 332 amino acid within the human SPRY domain’s primary restriction determination patch to encode glutamine (Q), rather than its current configuration of arginine (R). When they repeated the infection process of CrFK cells expressing the altered human TRIM5?, they found that the R332Q mutation did more than just decrease its efficiency at restricting PtERV1—they found that it more than doubled its efficiently at restricting HIV1.

    Since PtERV1 is extinct, it no longer exists in the environment to do humans any harm, thus the ability to restrict it is neither beneficial nor detrimental—it’s neutral. This renders the loss of efficient PtERV1 restriction likewise neutral.

    HIV1, on the other hand, does still exist in the environment, and does humans quite a bit of harm. Thus, the R332Q mutation causes an increase of beneficial ability with no loss of beneficial ability, rendering it a beneficial mutation.

    So now let’s back up a bit, and get to the details of the first step; establishing common ancestry. The following is a sampling of some vestigial evidence, as well as some of the strongest evidence; that of the genetic variety:

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    Evidence for the model of common ancestry:

    Vestigial evidence (subset of morphological evidence):

    I. Introduction

    II. Vestigial structures
    1. eyes of the blind marsupial mole

    III. Vestigial anatomical configurations
    1. paths of the left recurrent laryngeal nerve
    2. path of the phrenic nerve
    3. embryological placement of mammalian gonads

    IV. Vestigial reflexes
    1. emotive and thermoregulatory contraction of human Arrectores pilorum
    2. hiccupping (homologous to amphibian water gulping)

    Genetic evidence:

    I. Endogenous retroviruses
    1st layer) the presence of ERVs in orthologous loci among species of various degrees of taxonomic separation, and of the nested hierarchies they fall into
    2nd layer) the comparative degrees of LTR-LTR discontinuity among orthologous full-length ERVs
    3rd layer) shared mutations among orthologous ERV and the identical nested hierarchies they fall into

    II. Pseudogenes
    1. L-gulono-gamma-lactone oxidase
    a. shared mutations and the nested hierarchies they fall into

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    Vestigial Evidence:

    ——————————————–

    Vestiges are structures, anatomical configurations, and reflexes that are most parsimoniously explained as remnants of common ancestors in the evolutionary history of a given organism. As such, they are one of many sources of evidence for the model common ancestry, on which the evolutionary model is based.

    The most common example of a vestige is the presents of small, internal hind leg bones in modern whales, passed on to them by their land-dwelling mammalian ancestors. The response by many creationists—such as Ken Ham of Answers in Genesis, or Kent Hovind, in his series, Lies in the Textbooks—is that if a proposed vestige has any function at all, it isn’t really a vestige, and that this applies to whales since their leg bones play a roll in reproduction. What these creationists fail to understand is that the definition of a vestige has never been an evolutionary remnant with no function—it’s always been an evolutionary remnant that has lost or nearly lost its original primary function. If it has any current function, it’s either a persisting secondary function, or a function gained sometime after the lost of its primary one—a process called exaptation, or co-opting.

    Understanding exaptation and how it occurs is central to understanding why the creationist objection fails to address the issue of vestigiality, so let’s take a look at the analogy given in the second figure (on page 362) in the publication entitled “The Evolution of Complex Organs” by Dr. T. Ryan Gregory. The description reads:

    A simple example of exaptation and secondary adaptation. A The original and still primary adaptive function of coins is as currency. B A coin co-opted into a new exaptive role as an instant lottery ticket scraper. Coins would always have been capable of scraping tickets, but this function did not become apparent until an environment arose in which instant lottery tickets were abundant. Though functional as scrapers, coins are somewhat difficult to hold and may not reliably be on hand when needed. C A secondary adaptation that enhances the novel function of a coin as a ticket scraper by incorporating it into a keychain that is easier to grip (US Patent #6009590, “Lottery ticket scraper incorporating coin” by K.M. Stanford 2000). In this case, a second preexisting structure (key ring) was co-opted into a function as a carrier for a lottery ticket scraper (Gregory, 2008, p.362).

    For an explanation of six ways exaptation can occur—with examples of each—refer to pages 361 through 363 of Dr. Gregory’s publication on complex organ evolution. And as Dr. Gregory notes on page 363:

    The important point regarding exaptations, then, is that the current function of a feature may not reflect the reasons for its origin. Rather, the feature may only have come to occupy its current role comparatively recently.

    Now that it’s been established what vestiges are, how they form, how they provide evidence for common ancestry, and how the creationist objection from functionality is invalid, let’s look at specific examples of structural, anatomical, and reflexive vestiges.

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    II. Vestigial structures:

    1. Unlike most animals with eyes, “the marsupial mole (Notoryctes) is blind and has vestigial eyes that are hidden under the skin. Furthermore, the lens and pupil are absent, and the optic nerve is reduced (Springer et al., 1997, p.13758).” The deterioration of Notoryctes’ eyes are also accompanied by the deterioration of the interphotoreceptor retinoid binding protein gene; a genetic vestige.

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    III. Vestigial anatomical configurations:

    1. The left recurrent laryngeal nerve (RLN) branches from the left vagus nerve near the heart, and the right RLN braches off a bit further up from the right vagus nerve. The left RLN loops under the ductus arteriosus, and right RLN loops under the right subclavian artery. They both then travel up into the neck to innervate the larynx and its surrounding muscles. Both of these recurrent paths are quite inefficient in mammals; adding over a foot of unnecessary length to the human left RLN. This is especially true in giraffes, where the added length is absurd—more than fourteen signal-slowing, energy inefficient feet are added to the left RLN alone. This data doesn’t fit the model of creation, as it would be easier and more beneficial to simply branch the RLNs off higher on the vagus nerve and avoid arterial looping completely. The evolutionary model, however, is a precise fit; especially when one considers fish anatomy. In fish, several branches extend from the vagus nerve, each looping around arterial arches that connect the dorsal and ventral aorta between each gill slit (Ridley, 2004, p.281-282; Berry & Hallam, 1989, p.83). This is powerful evidence that mammals and fish share ancestry, that the RLNs and ductus arteriosus are remnants of the vagus nerve branches and sixth arterial arch in those ancestors, and that the configuration in mammals is a vestige resulting from decent with modification.

    2. The gonads of sharks, other fish, and even humans develop in same place—the chest. This works well for sharks, since they stay there, but in human males, the gonads need to travel all the way down into the scrotum to keep cool. This causes an unnecessary looping of the spermatic cord, which causes a weakness in there body wall, leaving them prone to developing a hernia (Shubin, 2009, p.64-66). This is consistent with decent, with modification, from an ancestor we share with modern fish.

    3. The path of the human phrenic nerves begins at the base of the skull, and goes through the body cavity to the diaphragm. This is an efficient path to amphibians’ gills, which are in the neck, but is an inefficient path to the diaphragm, in humans. The irritation of these nerves—made likely by their placement—can cause problems with breathing, including hiccups (Shubin, 2009, p.66-67); a reflexive vestige.

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    IV. Vestigial reflexes:

    1. Signals are sent from the human brain, through the phrenic nerves, that induce synchronous spasms of the diaphragm. The subsequent sharp inhalation of air closes the epiglottis. This is commonly known as the hiccups. The evidence, as laid out by Dr. Straus (2003), that it is a vestigial reflex is as follows:

    a) Amphibians have homologues motor pathways that cause similar inhalation (of water into their gills) and closing of the epiglottis (to prevent aspirating water into their lungs).

    b) Motor pathways necessary for hiccupping complete embryological development before those of lung functionality do.

    c) Both hiccups and amphibian water gulping are inhibited by the detection of high CO2 blood concentration by chemoreceptors.

    d) Both hiccups and amphibian water gulping are outright stopped by the binding of (RS)-4-amino-3-(4-chlorophenyl) butanoic acid (Baclofen) to gamma-aminobutyric acid B receptors.

    2. In many particularly hairy vertebrates, such as apes, dogs, and rats, the involuntary erection of hairs via the contraction of muscles, called Arrectores pilorum, act as an efficient means of thermoregulation. This is achieved by trapping insulating air against the skin. Humans, however, are not hairy enough for this reflex to have any significant effect in regulating their temperature. The involuntary erection of hair is also an emotive reflex in many vertebrates, increasing their apparent size to intimidate predators when afraid, to intimidate rivals when angry, or to convey various other emotions (Darwin, 1872, p.95-102). The only secondary purpose Arrectores pilorum serve is in aiding the sebaceous gland in secreting sebum (Song et. al., 2007).

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    Genetic Evidence:

    ——————————————–

    I. Endogenous Retrovirus:

    A powerful source of evidence that modern species diverged from ancestral species via descent with modification is that of endogenous retroviruses (ERVs). ERV evidence consists of three independent layers that corroborate one another. As will be further discussed, the three layers of ERV evidence are: 1) the sharing of ERVs in orthologous loci among organisms of varying degrees of taxonomic separation, and the nested hierarchies that these shared ERVs are arranged in; 2) the examination of shared mutagenic discrepancies between shared ERVs, so as to infer relative sequence of insertion; and 3) the nested hierarchies of shared mutations among given orthologous ERV’s. But before this evidence can be examined, one must have a firm understanding of retroviruses and how they infect cells.

    In order to replicate itself, a retrovirus needs to use the molecular machinery of a host, and it begins the process by first binding its extracellular and transmembrane glycoproteins to a cell’s coreceptors. The capsid—containing the retroviral enzymes reverse transcriptase, integrase, and protease, as well as two copies of the retroviral genome—is inserted into the cell’s cytoplasm, where it uncoats. Unlike a virus, with its genome comprised of DNA, now in its host cell, a retrovirus reverse-transcribes its genome from RNA to DNA via reverse transcriptase. Protease then process the viral DNA by removing a dinucleotide off each 3′ end, and integrase inserts it in the host cell’s genome (Targeting HIV replication, n.d.). Once integrated, and in DNA form, its genome is known as a prototype retrovirus, or provirus. Upon integration, the cell is allowed to divide, and eventually the presence of certain environmental conditions trigger proviral activation. Copy after copy of the retrovirus is produced as virions bud off, mature, and go on to infect other cells, leading to the death of the infected cell.

    The would-be problem for the retrovirus is that once the cell replicates and activation occurs, the provirus relies on the RNA polymerase II of the host cell to transcribe it into viral RNA to be packed into its virions. Unfortunately, RNA polymerase II is designed to transcribe messenger RNA (mRNA) for translation into polypeptides, so although it uses promoters to initiate transcription, since they don’t code for amino acids, it doesn’t transcribe them. With only one set of promoters on the proviruses, the newly transcribed retroviral genomes would have no promoters at all. When the virions that contained them would reverse transcribe them and insert them into new host cells, the second round of activation and transcription would never be able to occur.

    Retroviruses circumvent the problem of vanishing promoters by simply polymerizing copies of them during reverse transcription. They achieve this by possessing identical sections of DNA, called repeats, on either terminus of their genome. Early in the process of reverse transcription, the first jump occurs, in which the transfer RNA (tRNA) primer detaches and the DNA repeat hybridizes with the remaining RNA repeat at the genome’s 3′ terminus (Cann, n.d.). Given the relatively small size of the repeats, if they are not identical, they cannot hybridize. As for the 5′ unique (U5) and 3′ unique (U3) sections, a copy of each is polymerized on the opposite terminus. Between the necessity of identical repeats, and the duplication of unique sections, the resulting U3-R-U5 sections, called long terminal repeats (LTRs), must likewise be identical at the time of insertion. This will become very important later on in examining the second and third layers of ERV evidence.

    Once the provirus has been processed, integrase begins the process of insertion by breaking two of the host genome’s phosphodiester bonds between a 3′-hydroxyl group and 5′-phoshphate group, allowing the provirus to insert in a highly randomized manor (Skinner et.al., 2001). Different retroviral integrases have minor statistical biases for insertion within the general area of “chromosomal regions rich in expressed genes… CpG islands… active genes… [and areas] near transcription start sites (Mitchell et al., 2004),” but these biases are so minor, that they require thousands of trials (3127 used in the Mitchell study) just to detect them. For the most part, the process of integration is observed to be quite random.

    The two phosphodiester bonds integrase breaks are on either strand of the host DNA. Rather than breaking the bonds between the two nucleotides of a single base pair, integrase separates the breaks by several base pairs, forming a jagged cut (Skinner et. al., 2001). Since each of the two strands of DNA are complimentary to one another, the overhangs on either end of the jagged cut are likewise complementary. When nucleotides are polymerized to fill the gaps, the end result is two identical stretches of the DNA between the two initial breaks flanking the inserted provirus. This effect is called target site duplication. The second effect of the insertion is the displacement of DNA. This can be observed by comparing orthologous genomic sections in species with and without the provirus. The ones with the provirus display target site duplication, where as the ones without the provirus display a continuous stretch of DNA with a single copy of the duplication as a portion of it (Polavarapu, Bowen, & McDonald, 2006). The presents of these two effects ensures that you are dealing with an insertion.

    The targets of retroviruses are usually somatic cells, but if the infected cell happens to be a sperm or egg cell, known as a gamete, or a testicular or ovarian cell that divides into a gamete, that gamete may be used to produce an offspring. In such a case, the provirus becomes a permanent fixture within the offspring’s genome. Its permanence is due to fact that “there is no mechanism for removing proviruses precisely from the genome, without leaving behind a solo LTR or deleting chromosomal DNA (Johnson and Coffin, 1999).” Although the retrovirus was foreign to the organism it infected, and thus would be considered exogenous to that organism, once passed on to the organisms offspring, it would be present in the offspring’s natural, healthy state, and thus would considered endogenous to it.

    Now being passed on from one generation to the next, ERVs accumulate copying mistakes by DNA polymerase during subsequent host cell replication. Since ERVs are generally not conserved, they accumulate mutations at the same rate as introns. And, as with introns, over time, the mutations can become fixed in the host population’s gene pool (Boeke and Stoye, 1997. In Coffin, Hughes, & Varmus, 1997). Given enough time, enough mutations accumulate to render the ERV incapable of activation.

    If two or more individuals have ERVs of the same family at the same loci, one might think that there exist two plausible explanations: 1) that a retrovirus inserted in a common ancestor, and was passed on to the individuals via sexual reproduction; or 2) that separate retroviruses inserted in the same loci in separate ancestors, and were passed on to the individuals from each respective ancestor. What rules out the latter of these explanations is the highly random nature of integration discussed earlier, and shared mutation among orthologous ERVs, to be discussed shortly.

    Thus, we can then conclude—regardless of how many individuals have a given retrovirus in the same locus of their genome—that retrovirus necessarily inserted within the genome of a cell of a single ancestral organism common to each of them, and was passed on to those individuals via sexual reproduction. If the organisms are of the same species, the shared ERVs are referred to as paralogous, and the individual from which the insertion originated may likely have also been of the same species. If, however, the organisms are of different species, the shared ERVs are referred to as orthologous, and that is when things get interesting.

    Layer 1

    When we examine the collective genome of Homo sapiens, we find that a portion of it consists of ERVs (IHGS Consortium, 2001). We also find that humans share some of them with Chimpanzees, as well as the other Hominids (great apes) (Johnson and Coffin, 1999), Cercopithecids (old world monkeys) (Lebedev et. al., 2000), Platyrrhins (new world monkeys) (Steinhuber et. al., 1995), and even Prosimians (Anderssen et. al., 1997). Since humans don’t and/or can’t regularly procreate and have fertile offspring with members of these species, and thus don’t make sizable contributions to their gene pools, and vice versa, their inheritance cannot have resulted from unions of modern species. As previously mentioned, parallel integration is ruled out by the highly random target selection of integrase. And even if it was far more target-specific than observed, it would require so many simultaneous insertion and endogenizations that the evolutionary model would still be tremendously more parsimonious. This leaves only one way an ERV could have been inherited: via sexual reproduction of organisms of a species that later diverged into the one the organisms that share the ERV belong to, i.e. an ancestral species—simply put, humans and the other primates must share common ancestry.

    Not only are there many ERVs shared among primates, but they are shared in hierarchical subsets of the whole. Each set falls within another set, giving an unbroken line of inheritance for every species (Johnson and Coffin, 1999; Lebedev et. al., 2000; Steinhuber et. al., 1995; Anderssen et. al., 1997). This pattern is called a nested hierarchy. These patterns further corroborate that the many species of primates share common ancestry, and necessitate a specific sequence of divergence from one ancestral species to the next. They are wholly inexplicable by the model of uncommon ancestry.

    Layer 2

    As previously explained, although the LTRs of a provirus must be identical upon insertion, once endogenized, they begin accumulating mutations. Any mutations to one LTR become quite apparent, as they are not accompanied by the same mutations in the other. Thus each mutation causes the ratio of discontinuity between the two LTRs of a full-length ERV to increase. Since ERVs in orthologous loci among greater numbers species of wider taxonomic separation correlate to older insertions, if the evolutionary model is correct, they should also have higher ratios of discontinuity between their LTRs. And what do we find? We find just that; a pattern, where the degree of a shared ERVs’ LTR-LTR discontinuity is proportional to the degree of taxonomic separation between the species that share it (Johnson and Coffin, 1999). There is deviation from the pattern—particularly in the gorilla lineage—likely caused by viral transfer and interelement recombination/conversion (Hughes & Coffin, 2005) and viral transfer (Belshaw et. al., 2004) —but the pattern is holds for many full-length ERVs and is explainable only by decent with modification from a specific series of common ancestral species.

    Layer 3

    When the mutations in shared ERVs are examined, many are found to be identical to others. Just as will the distribution of ERVs, some shared mutations within a single shared ERV fall into nested hierarchies; some are shared by all, many by subsets of the whole, and each set falls within another set (Johnson and Coffin, 1999). Despite deviation caused by the same mechanisms effecting LTR-LTR discontinuity ratios, some of these nested hierarchies of mutation match those of distribution. Part of what makes this such powerful evidence for the evolutionary model is that is that ERV distribution and mutation rely on entirely different mechanisms; the function of integrase and the DNA replication complex, respectively. That the two nested hierarchies match at all is only explicable by common ancestry.

    In summary, the three layers of ERV evidence that have just been laid out are as follows:

    Layer 1: the presence of ERVs in orthologous loci among species of various degrees of taxonomic separation, and of the nested hierarchies they fall into.

    Since they’re passed on through sexual reproduction, ERVs fixed in orthologous loci in different species necessitates the past presence of a species ancestral to both, that has since diverged into the two modern ones. And the patterns of their distribution indicate a specific sequence of divergence.

    Layer 2: the comparative degrees of LTR-LTR discontinuity among orthologous full-length ERVs.

    Since LTRs are identical upon reverse transcription and subsequent insertion, greater divergence correlates to an older insertion. Thus the patterns of discontinuity indicate sequences of divergences consistent with those indicated by distribution.

    Layer 3: shared mutations among orthologous ERV and the identical nested hierarchies they fall into.

    Since mutations accumulate and fix in populations of organisms, the distribution of shared mutation indicate a sequence of speciation events consistent with that which is indicted by both distribution and LTR-LTR discontinuity.

    There are three common responses to ERV evidence by Creationists; the first is to ignore most of it—most notably of which being the patterns of distribution and mutation.

    The second is to use deviation from these patterns they fail to address as justification for dismissing ERVs outright. For example, they ignore viral transfer and interelement recombination/conversion (Polavarapu, Bowen, & McDonald, 2006) and state that the few lineage-specific PtERVs with uncharacteristically high LTR-LTR divergence ratios completely invalidates the second layer of evidence. What they conveniently overlook is that such deviation is to be expected, given the complexity of biological systems, and that it is the patterns they ignore that provide some of the strongest evidence for the evolutionary model.

    The third common response is to use red herrings to dismiss ERV evidence, such as that of functionality. There is no question that some ERVs have functions in organisms, but there are no wholly functional ERVs—only functional components, with the remainder deleted or mutated into non-functionality.

    For instance, the contribution of enJS56A1 and enJS5F16 (of the mere ~20 enJSRVs) to placental growth/differentiation regulation is achieved solely by their env genes with open reading frames (Dunlap et al., 2006; Palmarini et al., 2000). Although they also have an open gag reading frame (causing gag-gag interaction that restricts pathogenic JSRVs), they are the only ones known to have this (Mura et al., 2004). And every studied enJSRV has a closed pol reading frame (Murcia, Arnaud, & Palmarini, 2007).

    Another example is the transcriptional contribution of LTRs to genes’ promoters:

    1) Not only are most ERVs not at a loci that even makes it possible for them to contribute to transcriptional activity, but most ERVs have recombined into solo LTRs. Since only the LTRs of active full-length ERVs can contribute (Cohen, Lock, & Mager, 2009, p.107), even most ERVs in the right position have no effect. Just as with enJSRVs, these ERVs represent a very small percentage of the whole.

    2) The actual genes of these ERVs contribute nothing—only their promoter sequence-rich LTRs do. Again, just as with enJSRVs, these are examples of functional ERV components, rather than functional ERVs.

    It is the same with every case observed; again, there are no “functional ERVs;” only a small percentage of ERVs with functional components.

    But it’s a moot point, because we know that ERVs are insertions:

    The hallmark of an insertion is a displacement of chromosomal DNA, and the hallmark of insertion by integrase is the presents of target site duplication, due to the way it attacks the 5′ and 3′ phosphodiester bonds with an offset of a few base pairs (Skinner et al., 2001). Since ERVs are accompanied by target site duplications and DNA displacement, they are necessarily endogenized/fixed proviral insertions.

    So any functional components are necessarily post-insertion exaptations, and the fact that they are necessarily insertion means that they can not be part of any ‘original design.’ The issue of functionality is simply a red herring, when discussing how ERVs necessitate common ancestry.

    Again, for an understanding of exaptation (p.361-363), including various “paths to exaptation,” refer to the first half (p.358-366) of “The Evolution of Complex Organs” by Dr. Gregory (2008).

    Ultimately, the best way to respond to such claims—after having addressed their points specifically, of course—is to relentlessly drive home what they seem least willing to discuss; that deviation from patterns is to be expected, and that the corroboratory patterns of distribution and mutation are solely explicable by the evolutionary model.

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    II. Pseudogenes:

    A pseudogene is a former gene that has mutated to the point of inactivation. By comparing the polymorphism of psuedogenes in orthologues loci in other species, any nested hierarchies they fall into can be identified. This is aided by the identification of an orthologous active gene in other species, which the psuedogene is a deactivated copy of. The following is an example of how a pseudogene can be evidence of common ancestry:

    L-gulono-?-lactone oxidase (GULO) is the forth and final enzyme of the metabolic pathway that converts glucose in to L-ascorbic acid (vitamin C); a critical vitamin in biosynthesis; acting as a cofactor, substrate, or electron donor for numerous enzymes. Strepsirrhines and most other plants and animals have GULO, giving them immunity to deadly vitamin C deficiency (scurvy). Haplorrhines (Inai, Ohta, & Nishikimi, 2003), the remaining primates, along with pigs (Hasan et. al., 1999), guinea pigs (Nishikimi, Kawai, & Yagi, 1992), and certain fish and birds are prone to vitamin C deficiency. When we look at the position on their genomes where the GULO gene is located in organisms that do produce the enzyme, we find a mutated version of it. The mutations render it incapable of making GULO, or even activating in the first place. This mutated, inactive version of a gene is an example of what is called a pseudogene; in this case, a GULO pseudogene (GULOP). The original inactivation and subsequent accumulation of the majority of observed mutations was likely the result of the introduction of a premature stop codon found in the same place in all haplorrhines. And that isn’t the only GULOP mutation shared by all haplorrhines—there are many of them, with many more shared by subsets of the whole (Ohta & Nishikimi, 1999). Each set falls within another set, giving an unbroken line of inherence for every species. This pattern is called a nested hierarchy. Since the shared mutations necessitate inheritance from ancestors common to each member of each species that shares them (common ancestral species), which have since diverged—with the nested hierarchy they fall into both corroborating this and necessitating a specific sequence of divergence—and since GULOP can no longer perform its primary function (if any) of producing GULO enzymes, GULOP is a vestige of these ancestors.

  2. Jon says:

    References:

    Belshaw, R., V. Pereira, A. Katzourakis, G. Talbot, J. Paces, A. Burt, and M. Tristem. “Long-term reinfection of the human genome by endogenous retroviruses.” Proceedings of the National Academy of Sciences USA 101.14 (2004): 4894-899.
    tinyurl(DOT)com/Belshaw2004

    Berry, R. J., and A. Hallam, eds. Encyclopedia of Animal Evolution. 1st ed. Facts on File, 1989.

    Boeke, J. D., and J. P. Stoye. “Retrotransposons, endogenous retroviruses and the evolution of retroelements.” (1997). In. Coffin, J. M., S. H. Hughes, and H. E. Varmus. Retroviruses. New York: Cold Spring Harbor Laboratory Press, 1997.
    tinyurl(DOT)com/CoffinRetroviruses

    Cann, Alan. “Retroviruses.” MicrobiologyBytes. Web. 26 Oct. 2009.
    tinyurl(DOT)com/Cann2009

    Cohen, C. J., W. M. Lock, and D. L. Mager. “Endogenous retroviral LTRs as promoters for human genes: a critical assessment.” Gene 448.2 (2009): 105-14.
    tinyurl(DOT)com/Cohen2009Dec15

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