Print Email Download

Paid Writing Services

More Free Content

Order Now

Instant Price

Search our Site


Post Mortem

Chapter I
INTRODUCTION

"Watson, can you determine cause and time of death?" I knelt over the woman and began a cursory examination... “Rigor mortis has set in, so I'd estimate she's been dead about 10 to 12 hours." Holmes stood up and brushed himself off with his hands. "So, that puts her death between midnight and 2 am”(Anonymous 2007).

After the question of cause of death; the question of time of death is the most sought after piece of information associated with a medical death investigation. As a consequence, death investigators find themselves in need of a means of ascertaining the period of time between when an individual's body is found and when they died, sometimes referred to as the post mortem interval. Establishing the time of death through the determination of post mortem interval may have a direct bearing on the legal questions of guilt or innocence by confirming that a suspect's alibi covers the period when the victim died, or demonstrating that it does not. If the time of death can be established to within hours, days, months or even years, an individual may be able to prove that they were at some other place at that time. On the other hand, if the suspect is known to have been in the vicinity of the victim during the appropriate time period, then they can be shown to have had an opportunity to commit the crime.

Currently, there are multiple techniques for determining post mortem interval that incorporate methods in almost every discipline of forensic science. Depending on the circumstances, these techniques can yield results that vary from a narrow accurate estimate (video of the victim, the victim's stopped watch etc.) to a wide range estimate (counting tree rings on trees growing over or through the remains). Regardless of the of the method used, the calculation of post mortem interval is at best an estimate and should not be accepted as accurate without considering all of the factors that can potentially impact the result.

Post Mortem Interval Estimation

“For everything there is a season,
And a time for every matter under heaven:
A time to be born, and a time to die…” Ecclesiastes 3:1-2

The techniques currently utilized for estimating post mortem interval can be broken down into two broad categories based upon the methodology used. The first of these categories are the concurrence-based methodologies. Concurrence based methods relate or compare the occurrence of a known event, which took place at a known time, with the occurrence of death, which took place at an unknown time. Examples of concurrence-based methods include the determining the years of manufacture of clothing found on a body, tree ring development, dates on personal effects, etc. Concurrence based methods rely on both evidence associated with the body, and anamnestic evidence such as the deceased's normal pattern of movements. The second grouping of techniques include rate of change methodologies. Rate of change-based methodologies measure some aspect of a evidence, directly associated with the body, that changes at a known or predictable rate and is started or stopped at the time of death. Examples of the rate of change based methods include body temperature, tissue decomposition, insect succession and bone weathering. Some of these methodologies can be considered to fall into both categories. Examples of these would be tree ring development (Coyle, Lee et al. 2005) and insect succession.

Previous post mortem interval Estimation Methods

The variety of approaches for estimating post mortem interval spring from the varied expertise and experiences of their proponents as such the different methods tend to be focused on the immediate needs of the investigator, and limited to a particular stage of the post mortem interval or type of observation. As a consequence, the period of time for which a procedure is effective will overlap others.

Algor, Rigor and Liver Mortis

“Tis after death that we measure men.” James Barron Hope

The earliest recorded methods for estimating early post mortem interval were a rate of change methodology based on the most easily observed changes. The cooling of the body after death (algor mortis), the gradual stiffening of the body (rigor mortis) and the fixed pooling of the blood resulting in discoloration of the lower portions of the body (livor mortis) can be easily assessed with minimal or in some instances no instrumentation. Since the time of the ancient Greeks when the following rule of thumb was developed:

Warm and not stiff: Not dead more than three hours;

Warm and stiff: Dead between 3 and 8 hours;

Cold and stiff: Dead between 8 and 36 hours;

Cold and not stiff: Dead more than 36 hours; (Starkeby 2004)

until modern times, the basis of most temperature based post mortem interval analyses is the assumption that the human body, which averages 98.2 oF +/- 1.3 oF (Mall and Eisenmenger 2005), was at 98.6 oF (Mackowiak, Wasserman et al. 1992) at death and that after death the body looses heat in a predictable manner.

There have been many temperature based methods for estimating post mortem interval. As early as the 1800s, Dr. John Davy had developed a method using the fall in body temperature (algor mortis), measured rectally, to determine the post mortem interval (Henssge and Knight 2002). This method was refined by De Saram by recording detailed temperature measurements collected from executed prisoners (De Saram G. 1955). More recent approaches to this technique have included measuring rectal temperature, body surface temperature, ear canal temperature, eye socket temperature and liver temperature (Simonsen, Voigt et al. 1977; Henssge and Knight 1995; Baccino, De Saint Martin et al. 1996; Kanetake, Kanawaku et al. 2006).

Improvements to these techniques have included multiple progressive sampling, and the introduction of concepts such as the initial temperature plateau, core temperature, heat gradients, the effects of insulation, the ratio of surface area to volume, the effects of humidity and the effect of conductive surfaces, Microclimates and postmortem skin cooling (Green and Wright 1985; Nokes, Flint et al. 1992; Nelson 2000).

However, most methods that attempt to use body temperature changes to determine the post mortem interval are hampered, as most methods are, by individual variability. Even when complex calculations and algorithms have been designed to model for tissue density, initial temperature distribution, post mortem exothermic reactions and heat loss, these refinements have not appreciably narrowed the estimate window for post mortem interval. Multiple studies outlining instances of initial temperature increase of a body soon after death (Hutchins 1985) associated with post mortem chemical changes such as rigor mortis, cell lysis and the conversion of cellular energy production to anaerobic respiration (Nelson 2000); variations in the core body temperature ranging from 0.5 - 1.2 °C during a 24 hour period (Chisholm 1911; Mackowiak, Wasserman et al. 1992); the effect of variable environmental temperatures (Green and Wright 1985; Green and Wright 1985); and the effect of environmental temperature on overall body surface temperatures (Mall, Hubig et al. 2002) have all contributed to limit the usefulness temperature as a consistent indicator of post mortem interval. Additionally, once the body has reached ambient temperature temperature ceases to be a factor. Marshall said it best when he said ‘‘It would seem that the timing of death by means of temperature can never be more than an approximation''(Henssge and Knight 1995).

Soft and Hard Tissue Decomposition

“Now, a corpse, poor thing, is an untouchable and the process of decay is, of all pieces of bad manners, the vulgarest imaginable…” Aldous Huxley

Cadaveric decomposition is a complex process that begins immediately following death and proceeds beyond the time when recognizable human remains have ceased to exist. Decomposition can be broken down into two major stages. The first stage, soft-tissue decomposition, is caused by autolysis and putrefaction. Autolysis is the digestion of tissue by cellular enzymes and digestive processes normally present in the organism. Putrefaction is the digestion of whole tissues systems caused by the enzymatic activity of fungi and bacteria that are either present in the organism or the environment that opportunistically invade the tissue. Both autolysis and the microorganisms responsible for putrefaction are normally held in check in living organisms. However, when an organism dies the cellular and systemic mechanisms responsible for regulating autolysis and inhibiting putrefying microorganisms stop. “Without these controlling processes the body becomes fancy (bacterial) culture media” (Carayannopoulos 1992). These early postmortem changes in soft tissues can be used to provide an estimate of the post mortem interval from death until skeletonization. However, the rate of soft tissue decomposition can be dramatically affected by both internal and external factors that affect the body (i.e. ambient temperature, cause of death, scavenging, trauma, environmental conditions, clothing, body size, mummification and adipocere formation) (Rodriguez and Bass 1985; Micozzi 1986; Mant 1987; Vass, Bass et al. 1992; Komar 1998; Campobasso, Di Vella et al. 2001). There are reported instances of rapid decomposition associated with acute illness (Frisch 2001) and the author is personally aware of an instance of a post mortem interval of less than eleven days resulting in complete skeletalization of an individual that died of complications related to Acquired Immunodeficiency Syndrome (Watson 1994). Additionally, there are a number of examples of bodies remaining intact for years after death (Bass and Jefferson 2003).

Beyond gross observation for assessing decomposition, researchers have developed multiple morphometric and chemical methods for assessing soft tissue decomposition. These have ranged from early (ca.1800s) methods such as the Brouardel method which examined the shift in flammability of putrefaction gases in the early post-mortem interval, and the Westernhoffer-Rocha-Valverde method examining the formation of crystals in the blood formed after the third day of putrefaction (Cengage 2006); to more modern methods such as ultrasound assessments of organ condition (Uchigasaki, Oesterhelweg et al. 2004) and the use of electron microscopy to examine measurable physical changes in mitochondria (Munoz, de Almeida et al. 1999) and platelet count (Thomsen, Kaatsch et al. 1999). Chemical methods used to assess time since death include the assessment of volatile organic compound formation (Vass, Bass et al. 1992; Statheropoulos, Spiliopoulou et al. 2005; Statheropoulos, Agapiou et al. 2007; Dekeirsschieter, Verheggen et al. 2009); the concentrations of non-protein nitrogen (Sasaki, Tsunenari et al. 1983; Gallois-Montbrun, Barres et al. 1988) and creatinine (Gallois-Montbrun, Barres et al. 1988; Brion, Marc et al. 1991).

Bony tissue decomposition, the second major stage of decomposition, consists of a combination of surface weathering due to environmental conditions (temperature, humidity, sunlight) and erosion from soil conditions (pH, mineral content, etc.) (Behrensmeyer 1978; Janjua and Rogers 2008). While not much detailed study has been done on the environmental factors that affect bony tissue breakdown, it has been established that environmental factors such as pH, oxygenation, hydrology and soil flora and fauna can affect the long term stability of collagen (Garlick 1969; Henderson 1987; Bell, Skinner et al. 1996). Collagen, the primary protenatious component of bone, slowly hydrolyzes to peptides and then to amino acids leading to the breakdown of the collagen-mineral bonds which weakens the overall bone structure leaving it more susceptible to environmental weathering (Henderson 1987). By examining the effects of related changes (cracking, flaking, vacuole formation, UV-fluorescence of compact bone) the investigator can estimate the period of time a bone sample has been exposed to weathering (Yoshino, Kimijima et al. 1991; Bell, Skinner et al. 1996; Janjua and Rogers 2008; Wieberg and Wescott 2008). Current methods of assessing time since death using bone weathering rely heavily upon the experience of the investigator (Knight and Lauder 1969) and are limited to immediately post skeletalization to 10 to 100 years based on environmental conditions (Haglund and Sorg 1997).

As with the assessment of soft tissue decomposition for time since death, investigators examining bone decomposition have supplemented observational methods with quantifiable testing techniques that analyze changes that are not directly affected by the physical environment (Lundquist 1963). Radiocarbon dating of carbon-14 and strontium-90 have been used to group remains pre and post 1950 (Taylor, Suchey et al. 1989; Maclaughlin-Black, Herd et al. 1992). Neis suggested that, with further study of strontium-90 distributions, determination of times since death should be possible (Neis, Hille et al. 1999). Bradley suggested that measuring the distribution of 210Pb and 210Po in marrow and calcified bone could prove forensically significant (Bradley 1993). This work was built upon by Swift who evaluated using 210Pb and 210Po distribution in conjunction with trace element analysis to provide a meaningful estimate of the post-mortem interval (Swift 1998; Swift, Lauder et al. 2001). Maclaughlin demonstrated that chemical changes due to environment could measurably affect isotope levels (Maclaughlin-Black, Herd et al. 1992). In addition to radionucleotide studies, investigators have also measured the changes in both organic (amino acids, urea, proteins, DNA) and inorganic compounds (nitrogen, potassium, sulphur, phosphorous) in bone. (Jarvis 1997; Prieto-Castello, Hernandez del Rincon et al. 2007).

Stomach Contents/Rate of Digestion

“Govern well thy appetite, lest sin surprise thee, and her black attendant Death.” John Milton

The presence or absence of food in the stomach is often used as an indicator of post mortem interval. Its use as an indicator of post mortem interval is predicated on the assumption that under normal circumstances, the stomach digests and empties at a predictable rate taking from two to six hours to eliminate a full meal (Jaffe 1989). If a person had eaten a light meal the stomach would empty in about 1.5-2 hours. For a medium-sized meal the stomach would be expected to take about three to four hours to empty. Finally, a large meal would take about four to six hours to exit the stomach. Regardless, it would take from six to eight hours for the initial portion of the meal to reach the large intestine (Hallcox 2007). This information, coupled with reliable ante-mortem information relating to when an individual last ate is used by some pathologists when providing an estimate of the times since death. It is for this reason, among others, that comprehensive autopsies usually include an examination of the stomach contents (Batten 1995; Siegel 2006).

Although it provides another useful indicator of time since death, there are serious limitations to the assessment of the stomach contents as an accurate indicator of time since death. Its reliance on reliable anamnestic evidence such as eating habits, the extent to which the victim chews their food (Pera, Bucca et al. 2002), the physiological state of the victim (Troncon, Bennett et al. 1994; Jayaram, Bowen et al. 1997; Lipp, Schnedl et al. 1997; Phillips, Salman et al. 1997) and the state of mind of the victim (Jaffe 1989); as well as verifiable antemortem evidence such as what the last meal consisted of (protein vs. fiber vs. fat)(Dubois 1985; Tomlin, Brown et al. 1993), the amount of liquid consumed with the meal, alcohol consumption and the time when it was consumed limits its usefulness to a small number of cases (Jaffe 1989). These factors combined with evidence that digestion can continue after death (Koersve 1951) makes the estimation of post mortem interval using stomach contents difficult at best.

Insect Succession

“Buzzards gotta eat, same as worms.” Clint Eastwood from the Outlaw Josey Wales

Insect colonization of a body begins within hours of death and proceeds until remains cease to be a viable insect food source. Throughout this period, multiple waves of colonization by different insect species, as well as multiple generations of previously established species can exist. Forensic entomologists can use the waves of succession and generation time to estimate the postmortem interval based on the variety and stage of development of the insects, or insect remnants, present on the body (Archer and Elgar 2003). In addition to information regarding time since death, forensic entomology can provide useful information about the conditions to which the body was exposed. Most insects have a preference for specific conditions and habitats when colonizing a body and laying their eggs. Modifications to that optimal habitat can interrupt the expected insect colonization and succession. The presence of insects or insect larva that would typically be found on bodies colonized indoors or in shade on a body discovered outside in direct sunlight may indicate that the body was moved after death (Sharanowski, Walker et al. 2008). Aquatic insects found on bodies discovered on land could indicate the body was originally in water (Wallace, Merritt et al. 2008; Proctor 2009).

Although insect succession varies by season, geographical location and local environmental conditions, it is commonly assumed to follow a predictable sequence within a defined habitat. While there are a multitude of studies that have examined regional succession patterns (Archer and Elgar 2003; Tabor, Brewster et al. 2004; Tabor, Fell et al. 2005; Martinez, Duque et al. 2007; Eberhardt and Elliot 2008; Sharanowski, Walker et al. 2008) these studies use different approaches towards defining habitat and assessing insect succession making cross-comparisons of their data difficult. Also, the majority of these studies do not rigorously address the statistical predictability of a species occurrence making their results of limited use as post mortem interval indicators (Michaud and Moreau 2009). Additionally, beyond the presence or absence of clothing, the majority of the post mortem entomological studies conducted do not examine non-habitat external factors that may affect succession. For example, only a few studies have been conducted that assess the affect of drug ingestion (George, Archer et al. 2009) or the presence of chemicals (bleach, lye, acid etc.) used to cover-up evidence (Charabidze, Bourel et al. 2009) on the insect life cycle. As with other means of assessing time since death, more extensive studies with different insect species and drugs in a wider variety of habitats is necessary.

Electrolyte Concentration

“Death is a low chemical trick played on everybody…” J.J. Furnas

Cellular activity does not immediately cease when an organism dies. Rather, individual cells will continue to function at varying metabolic rates until the loss of oxygen and metabolic substrates caused by the cessation of blood flow results in hypoxia (low oxygen). As cell metabolism shifts from aerobic to anaerobic, oxidative phosphorylation and ATP generation, the cellular processes keeping autolysis in check, begin to decrease and eventually cease all together. Without energy to maintain osmotic gradients membranes begin to fail. As lysosomal membranes begin to fail the enzymes within are released and begin consuming the cell from the inside out. With autolysis comes a cascade of metabolic chemicals, released ions, originally bound up in various cellular processes begin to diffuse due to the diffusion gradient according to Fick's law into the intracellular spaces (Madea 2005). Forensic researchers have used the presence, absence or effects of inorganic ions such as potassium, phosphorous, calcium, sodium and chloride as a means of estimating time since death (Schleyer and Sellier 1958). In most instances the higher the concentration gradient, the more suitable is the analyte for the estimation of the time since death. When analyzing body fluids for the purposes estimating post mortem interval, early researchers tended to focus their studies on body fluids such as, cerebrospinal fluid, blood and pericardial fluid (Schleyer and Brehmer 1958; Coe 1972; Henssge and Knight 1995; Yadav, Deshpande et al. 2007) with a few others examining other compartmentalized bodily fluids (Madea, Kreuser et al. 2001) and the largest numbers focusing on vitrious humor (Madea, Henssge et al. 1989; Ferslew, Hagardorn et al. 1998; Madea and Rodig 2006; Kumagai, Nakayashiki et al. 2007; Thierauf, Musshoff et al. 2009). Chemical methods used to assess these analytes in blood and spinal fluid as an indicator of post mortem interval have failed to gain general acceptance because, for the most part, they failed to produce precise, reliable, and rapid results as required by the forensic community (Lundquist 1963). Current chemical methods which have primarily focused on vitreous fluid tend to suffer from the same limitations demonstrated by the fact that with notable exceptions (Pounder 1995) very few statistically rigorous field studies on the reliability and precision of estimating post mortem interval are available in the literature (Coe 1993; Madea 2005).

Enzyme Activity

As previously discussed, cellular activity does not cease when clinical death occurs. In any circumstances where the cellular metabolism shifts from a homeostatic balanced state to an imbalanced state biochemical changes occur. Changes in the levels and/or activity of enzymes (i.e. cardiac troponin, c-reactive proteins, and G proteins) have long been used as indicators of cellular stress (Li, Greenwood et al. 1996; Katrukha, Bereznikova et al. 1998; Tsokos, Reichelt et al. 2001; Uhlin-Hansen 2001). Assessing similar changes in cellular biochemistry as a function of time since death provides investigators with a wide variety of tissues, testing methods and analytes for consideration. As a consequence, forensic investigators have assessed and suggested enzymes from heart, pancreas, muscle, blood and brain as potentially suitable markers for time since death (Wehner, Wehner et al. 1999; Wehner, Wehner et al. 2001; Kang, Kassam et al. 2003; Jia, Ekman et al. 2007; Poloz and O'Day 2009). Comparisons of total proteins analyzed ante and post mortem analyzed using two dimensional gel electrophoresis and Matrix Assisted Laser Desorption/Ionization Time-of-Flight have demonstrated changes in metabolic enzymes, (Jia, Ekman et al. 2007; Hunsucker, Solomon et al. 2008). Assessing the changes in enzyme activity provides examiners a means to assess time since death, in many instances long before visible cellular changes. However, in at least a few of these studies results indicate that enzyme degradation during extraction and partial enzyme activity observed with degradation products these markers better suited to qualitative analysis rather than quantitative analysis (Sabucedo and Furton 2003).

Muscle/Nerve Excitation

Both neurons and myocytes retain the ability to respond to electrical stimulation for at least a short period of time after organism death. (Sugioka, Sawai et al. 1995; Briskey, Kastenchmidt et al. 2002; Sams 2002). The response of nervous and muscle tissue to external electric stimulation has also been investigated and proposed as means to estimate time since death (Kline and Bechtel 1990; Straton, Busuttil et al. 1992).

Methods developed to investigate myocyte excitability assess the relative magnitude and duration of the muscle contraction during the application of external stimulation. To assess the contractile response, a combination of observational based assessments (Madea 1990; Jones, James et al. 1995) and measurement based assessments (Henssge, Lunkenheimer et al. 1984; Madea 1992) have been suggested and reported.

Similar investigations have examined post mortem excitation of nervous tissue by measuring a variety of neurological reactions to stimuli. These include the alteration of Compound Muscle Action Potential (Nokes, Daniel et al. 1991; Elmas, Baslo et al. 2001; Elmas, Baslo et al. 2002), lengthen of the refractory or non-propagating period immediately following the CMAP (McDowall, Lenihan et al. 1998), the extracellular impedance/resistance (Querido 2000), the chronaxie measurement or the time over which a current double that necessary to produce a contraction is applied before the contraction occurs (Straton, Busuttil et al. 1992) and the changes in the amplitude of the F-wave (the secondary CMAP observed after the initial CMAP) have all been examined, and been suggested as potential indicators of time since death.

The results of studies examining the response of excitable tissue to electric stimulation have been consistent in that the stimulation response varies predictably over time. However, suitability for absolute indicators of time since death remains in questions as investigators have reported contradictory results related to the effect of the manner of death on the stimulation response (Madea and Henssge 1990; Elmas, Baslo et al. 2002).

RNA Degradation

RNA degradation, both antemortem and postmortem, is a complex process that is not well understood. Unlike with DNA degradation, continuous degradation of inducible mRNAs by native ribonucleases is used as a means of translational control. After cell death these ribonucleases, no longer kept in check by the mechanisms of cellular homeostasis, combine with exogenous ribonucleases from bacteria and fungi to begin un-inhibited digestion of all cellular RNA. Investigators have noted extensive variability in RNA degradation rates in different tissues (Bauer 2007). Not surprisingly such variability appears to be related to the antemortem ribonuclease activity of the tissue; with relatively ribonuclease poor tissues such as brain and retina exhibiting greater RNA stability (Johnson, Morgan et al. 1986; Malik, Chen et al. 2003) when compared to ribonucleases rich tissues such as liver, stomach and pancreas (Humphreys-Beher, King et al. 1986; Finger, Mercer et al. 1987; Bauer, Gramlich et al. 2003). Additionally, but also not surprisingly, some constitutively expressed mRNAs have been shown to be more stable, or perhaps simply more prevalent, than inducible mRNAs (Inoue, Kimura et al. 2002). Additionally, while intrabrain mRNA levels are fairly constant, interbrain levels vary considerably (Preece, Virley et al. 2003). As a consequence of these observations, the degradation of RNA (total and/or mRNA) have been suggested as a potential analyte to assess time since death.

Researchers examining the effect of post mortem interval on RNA stability have examined a variety of targets (mRNA, both tissue specific and constitutively expressed, and total RNA) with an assortment of methods including Reverse Transcriptase (RT) PCR(Ohshima and Sato 1998; Fleige, Walf et al. 2006; Haller, Kanakapalli et al. 2006; Zhao, Zhu et al. 2006), RNA (cDNA) microarrays (Bahn, Augood et al. 2001; Catts, Catts et al. 2005; Son, Bilke et al. 2005; Popova, Mennerich et al. 2008) and quantitative RT-qPCR (VanGuilder, Vrana et al. 2008). Based on these studies, there are indications that beyond time and temperature, factors such as hypoxia, tissue pH, antemortem physiological conditions (coma, seizure activity and injury) postmortem transcriptional activity and RNA sequence can dramatically affect the stability and measurable levels of RNA (Burke, O'Malley et al. 1991; Harrison, Heath et al. 1995; Ohshima and Sato 1998; Catts, Catts et al. 2005; Bauer 2007). When examining the seminal question regarding time since death and temperature some researchers have reported temperature and time as significant factors affecting mRNA levels (Burke, O'Malley et al. 1991), while others have reported the reverse (Harrison, Heath et al. 1995; Preece and Cairns 2003). These contradictory data are not surprising given the changes in the specificity, sensitivity and application of the assays used; however, the ultimate question has not been resolved. What is clear from the research is that RNA degradation (mRNA or total) is a complex process (Preece and Cairns 2003; Preece, Virley et al. 2003; Heinrich, Lutz-Bonengel et al. 2007) effected by multiple factors indicating more study will be required before RNA degradation can be considered a reliable indicator of time since death.

DNA Degradation and its Effect on DNA Typing

Since the initial application of molecular biology techniques to samples of forensic significance in the latter half of the 1980s, forensic scientists have noted that increased exposure to environmental insults can negatively impact DNA quality. Developmental validation studies performed to evaluate the efficacy of new typing techniques (SWGDAM 2008) have found that environmental variables such as heat, high humidity, direct moisture, fungal/bacterial contamination and ultraviolet light can impact the quantity or quality of the DNA sample making them unsuitable for DNA analysis (McNally, Shaler et al. 1989; Graw, Weisser et al. 2000; Takayama, Nakamura et al. 2003; Bender, Farfan et al. 2004; Schneider, Bender et al. 2004; Niemcunowicz-Janica, Pepinski et al. 2007). During transitions in technology from Restriction Fragment Length Polymorphism (RFLP) analysis to Polymerase Chain Reaction (PCR) based testing, researchers noted that samples too degraded to produce an RFLP pattern could still produce profiles using a variety of PCR based markers that evaluated loci shorter in length (Hochmeister, Budowle et al. 1991). This finding supports the hypothesis that degradation in the forensic setting is (not surprisingly) processive. Additional research found that while the DNA in some samples like cadaveric blood and kidney tissue could degrade to the point where it was no longer suitable for DNA fingerprinting after as little as a week (Ludes, Pfitzinger et al. 1993); other samples such as bone (Hochmeister, Budowle et al. 1991; Frank and Llewellyn 1999) and teeth (Schwartz, Schwartz et al. 1991; Pfeiffer, Huhne et al. 1999) could, under most conditions, provide typeable DNA for months.

The fact that DNA degradation has a detrimental effect on larger genetic loci, and affects different tissues at different rates is considered to be of extraordinary forensic significance is evidenced by the numbers of studies that seek to examine, and overcome this effect (42 validation studies specifically mentioning DNA degradation from 1995-2009 in PubMed). This makes perfect sense when the observer considers the impact that degradation can have on selecting suitable samples and evaluating the resultant DNA profiles. However, a number of researchers have looked beyond the simple question of how degradation affects the typing of samples to broader questions such as the mechanisms of postmortem degradation (De María and Arruti 2004; Foran 2006) and synthesis (Oehmichen, Frasunek et al. 1988) and how that knowledge can be used to assist in the assessment of time since death.

DNA degradation by RFLP:

Since Sir Alec Jeffreys first applied Southern blotting (Southern 1975) techniques to the testing of forensically significant samples in 1985 (Jeffreys, Brookfield et al. 1985) DNA analysis has revolutionized forensic science. Restriction Fragment Length Polymorphism DNA analysis relies on variations in the lengths of DNA fragments generated by enzyme restriction. With restriction fragments ranging from approximately from 2 - 33 kilobases (Baird, Balazs et al. 1986) successful typing and analysis requires high quality (un-fragmented) DNA. Researchers noted from the outset that in some cases involving older and/or postmortem samples that DNA degradation, tied to the exposures of higher temperatures, resulted in the gradual disappearance of the longer fragments reducing the evidentiary value of older samples (Bar, Kratzer et al. 1988). At about the same time, researchers interested in assessing time since death were building on this work by comparing DNA degradation in rib bone from multiple individuals using Southern blotting and radioactive probing. Results of this early work indicated that variations in the rate of DNA degradation between individuals was not as significant as between samples exposed to different postmortem intervals and temperatures (Perry, Bass et al. 1988).

DNA Degradation by DNA Flow Cytometric Analysis:

Flow cytometry is a method that can be used to determine the relative nuclear DNA content in a cell. The flow cytometer analyses moving molecules in a suspension that are excited by a source of light (U.V. or laser) and in turn emit a signal. The signal is captured and converted into a graphical representation of the intensity of the fluorescence emitted. By analyzing the amount of fluorescence emitted, the analyst can evaluate the DNA content. The results of this measurement can be used to assess DNA fragment sizes from about 1-150 kilobases (Yan, Habbersett et al. 2000; Ferris, Habbersett et al. 2004).

Utilizing flow cytometry, researchers have investigated the effects of time related degradation/fragmentation on postmortem eukaryotic cells (Afanas'ev, Korol et al. 1986) with at least one investigator demonstrating that there is at least a loose correspondence between DNA degradation in “corpse” tissue and samples collected in vivo (Di Nunno, Costantinides et al. 1998). Forensic investigators have proposed using a variety of different tissues as a source of samples showing the best correlation between between DNA degradation and postmortem interval. Splenic cells have been recommended by several investigators (Cina 1994; Di Nunno, Costantinides et al. 2002; Boy, Bernitz et al. 2003). Other internal organs evaluated include blood and liver with hepatic tissue showing an almost linear correlation between the time since death and the level of DNA degradation (Di Nunno, Costantinides et al. 2002). “Non-organ” tissues suggested as suitable samples for flow cytometry evaluations of times since death include costicartilage cells and dental pulp (Boy, Bernitz et al. 2003; Long, Wang et al. 2005).

While investigators have found good correlations between time since death and DNA degradation measured using flow cytometry, limitations to the procedure as it was originally proposed have proven difficult to overcome for forensic samples. Flow cytometry requires a suspension of intact cells for staining and analysis. This makes solid tissue difficult to analyze without extensive manipulation. Additionally, the assay measures total DNA making the differentiation intact human DNA from bacterial and fungal DNA difficult. While several modifications to flow cytometry have made it more specific and sensitive (Goddard, Martin et al. 2006), they have not gained the same general acceptance as DNA amplification.

Post mortem degradation by single cell electrophoresis

Single Cell Gel Electrophoresis (SCGE), sometimes referred to as a comet assay, uses migration of DNA from cells encapsulated in agarose to measure the level of DNA fragmentation. A tissue sample is encapsulated in agarose and the DNA within the tissue is denatured. The DNA is electrophoresed through the encapsulating agarose and samples with degraded DNA generate smeared tails (hence the name comet assay). The stronger the signal from the tail, the more DNA damage present. Tail length is directly impacted by the size of the DNA fragments with more degraded DNA generating longer tails. The tail moment is the product of the fraction of DNA in the tail and the tail length and is related to the proportion of non-fragmented to fragmented DNA.

Forensic investigators have evaluated SCGE as a means of assessing time since death analyzing DNA from skeletal muscle, heart muscle, liver and kidney with increasing postmortem intervals (Johnson and Ferris 2002; Zhen, Zhang et al. 2006). Measurement of tail-length providing the strongest statistical correlation, based upon regression analysis. (Johnson and Ferris 2002). While a promising technology, SCGE lacks specificity being unable to differentiate between eukaryotic and prokaryotic DNA mixtures; is relatively insensitive requiring microgram quantities of DNA; and exhibits excessive interlaboratory variability due to a lack of consistent protocols and analysis techniques (Dhawan, Bajpayee et al. 2009). Despite these issues, several investigators have suggested SCGE as a useful quantitative assay to pair with RT-PCR (Alaeddini, Walsh et al. 2009). As with flow cytometry however, SCGE has not gained the same general acceptance that Polymerase Chain Reaction based assay's have.

DNA Degradation by in situ Labeling

Fluorescence In Situ Hybridization (FISH) is a technique used to detect specific DNA sequences on chromosomes. FISH uses a fluorescently labeled probe that is complementary to the DNA sequence of interest. Once bound, the probe can be detected using fluorescence microscopy or other fluorescence sensitive instrumentation.

Forensic FISH techniques using centromeric probes for sex chromosomes were developed to identify the sex of contributors to bloodstains (Pettenati, Rao et al. 1995) and for the selection and microdissection of male cells on slides with male/female cellular mixtures (Vandewoestyne, Van Hoofstat et al. 2009).

DNA Degradation Image Analysis/Staining

A variety of different nucleic acid staining techniques have been applied to the analysis of forensic evidence. As is often the case with technology, the use of nucleic acid staining as a means of assessing medico legally significant aspects of a case has been cyclical. Early applications of nucleic acid/chromatin staining such as the use of Kernechtrot-Picroindigocarmine (KPIC) staining , sometimes referred to as Christmas tree staining (Whitney 1897) and the Feulgen method with Schiff's reagent (Knight 1963) were first applied to the detection of sperm in sexual assaults. Modifications to these techniques further refined their efficacy in criminal cases allowing the determination of sex of hairs (Nagamori 1978). With the advent of DNA analysis for paternity (Jeffreys, Brookfield et al. 1985) and forensic testing (Gill, Jeffreys et al. 1985), nucleic acid staining moved from the forefront of forensic testing to a secondary position as an ancillary analysis technique used with RFLP (Waye and Fourney 1990; Laber, O'Connor et al. 1992) and PCR (Budowle, Chakraborty et al. 1991).

However, even as the majority of forensic scientists began moving away from staining nucleic acid as an analysis technique in and of itself, other researchers were developing new staining assays that had the potential to provide information useful to the forensic investigator. Researchers investigating DNA damage developed specialized staining techniques using fluorochromes that, under the proper conditions, interact preferentially with dsDNA allowing measurements of DNA denaturation/damage (Batel, Jaksic et al. 1999). Other researchers have developed new applications for the Feulgen method that combine staining with micro densitometry allowing increased sensitivity and accuracy (Hardie, Gregory et al. 2002). By applying these methods to a variety of different cell types (Lin, Liu et al. 2000; Chen and Cheng 2002) several investigators have reported both a linear relationship between DNA degradation and the early time since death (Lin, Liu et al. 2000; Shu, Liu et al. 2005; Chen, Yi et al. 2007; Liu, Shu et al. 2007) and a measurable effect of temperature the rate of degradation (Wang, Zhang et al. 2006). For the most part however, general staining lacks specificity being unable to differentiate between eukaryotic and prokaryotic DNA mixtures so requires intact tissue not exposed to bacteria making postmortem forensic samples unsuitable.

DNA Degradation by DNA Amplification

Polymerase Chain Reaction (PCR) (Saiki, Scharf et al. 1985; Mullis, Faloona et al. 1986) is a process used to copy or amplify, by several orders of magnitude, specific segments of a DNA fragment. PCR performs this amplification by reproducing in a test tube the normal process of DNA replication that occurs in the cell. Samples that start with only a few complete copies of template DNA can, with the aid of PCR, produce tens of millions of copies of the sequence of interest suitable for genetic analysis. This occurs because as PCR progresses, each replicated copy of the DNA produced by a previous copying cycle can then be replicated in the subsequent cycle(s), resulting in an exponential increase in DNA copies. PCR starts with an enzymatic reaction mix that contains an optimizing buffer, a thermostable DNA polymerase (often a variant of Taq polymerase), short DNA sequences (primers) complementary to the DNA target(s) of interest and dideoxyribonucleoside triphosphates (ddNTPs), the individual building blocks used to assemble new DNA strands). This reaction mix is then combined with the extracted template DNA and placed in a thermocycler. The thermocycler alternately heats and cools (cycles) the mixture allowing the replication process to occur with each cycle. The initial step in each cycle is denaturation, where the reaction mix is heated to near boiling and the double-stranded DNA template separates (melts) into two single strands. The reaction mix is then cooled to the optimal sequence dependent temperature for the primers to bind or anneal to the DNA template. During the extension step the reaction mix is heated to the optimal extension temperature for Taq polymerase to extend the primer making the new copy of the DNA molecule.

Since the mid 1980's, all DNA based methods for establishing human identity and/or kinship have relied on various methods of assessing length polymorphisms, sequence polymorphisms or both. As testing preferences shifted from large fragment analysis like RFLP (kilobase length) to shorter fragments such as VNTRs (several 100-1000 bases) to even shorter fragments like STRs (<100 - 350bases) there was a lessened need, and concomitant lessened focus on the condition/degradation level of the sample DNA. However, as PCR-STR analysis became more widespread and applied in more varied types of case work it became obvious that

Competitive PCR

Competitive PCR (c-PCR) is a quantitative PCR system that is based on the addition of a competitor DNA fragment with the same primer recognition sites as the target gene but producing a differently sized fragment (Zimmermann and Mannhalter 1996). Once amplified the fragments are separated using slab gel (agarose or polyacrylamide) electrophoresis or capillary electrophoresis. The signal from the target fragment and competitive fragment are compared to determine the quantity.

Researchers have used the competitive PCR process to develop a quantitative PCR assay that used probes targeting mitochondrial DNA hypervariable region I and the amelogenin locus. Using this assay, investigators were able to assess the changes in the quantity of DNA in tissues exposed to different conditions for different periods of time finding that DNA was well preserved in bone and fingernails but rapidly decreased in soft tissues regardless of the exposure time or conditions (Imaizumi, Miyasaka et al. 2004).

Real Time PCR

Real-Time quantitative PCR (RT-qPCR) is an amplification process that allows the quantification of the number of copies of a specific DNA sequence present in a sample as the sample is being amplified. The procedure relies on PCR combined with a means to continuously measure the increase in dsDNA PCR product. The initial publication describing quantitative PCR utilized the hybridization radioactively labeled probes to the amplified product followed by exposure to x-ray film (Kellogg, Sninsky et al. 1990). Subsequent researchers proposed modifications the using the 5'-3- exonuclease activity of Thermus aquaticus (Taq) DNA polymerase to release fluorescent dyes associated with binding probes and the exposure to film (Holland, Abramson et al. 1991) and capillary electrophoresis (Fasco, Treanor et al. 1995). However, while these assays were informative they could not be considered “real-time” in that they each required post PCR manipulation to produce data. The initial publication suggesting a real-time method proposed the use of intercalating dyes combined with a video to measure the increase in signal (Higuchi, Fockler et al. 1993). Subsequently, multiple researchers proposed adapting Holland's method for use with analytical thermocyclers that could measure the increase in fluorescence as the sample was amplified (Chiang, Song et al. 1996; Gibson, Heid et al. 1996; Heid, Stevens et al. 1996).

Over the last ten years RT-qPCR has become the standard method for forensic sample quantification (Green, Roinestad et al. 2005). The quantifiler human kit produced by Applied Biosystems targets a 62 base amplicon within an intron in the human telomerase reverse transcriptase gene (hTERT), chromosomal location 5p15.33 (NFSTC 2010). The process uses standard oligonucleotide primers for amplification, a minor groove binding TaqMan® reporter probe (Heid, Stevens et al. 1996) and the 5'-3' exonuclease activity to Taq polymerase described by Holland above. The reporter probe has a fluorescent tag on the 5'end, a fluorescent signal quencher on its 3' end and binds in the minor groove of one strand of the target DNA. During amplification as Taq extends the amplification primer bound to the strand with the reporter, the enzyme encounters and excises the 5'end of the reporter probe releasing the fluorescent tag. Once the fluorescent tag is no longer in proximity to the quencher it gives off a detectable signal when excided by UV light.

As the equipment necessary to perform RT-qPCR has become more prevalent in forensic laboratories; and forensic scientists have become more familiar with the procedure and results; forensic researchers began to recognize the potential for using RT-qPCR to assess both the quantity of DNA and the likelihood of successful genotyping. To achieve this researchers have proposed modifications to the procedure that would allow analysts to assess both total DNA and the degradation level of that DNA. The most straight forward modification was the incorporation of a longer amplicon (170-190 bp) to assess the level of degradation (Swango, Timken et al. 2006; Swango, Hudlow et al. 2007). The short amplicon included in the Quantifiler® kit (62bp) is much shorter than the amplification products generated during genotyping (106-350 bp) resulting in instances where sufficient quantities of DNA are present to type but no profile can be generated due to the quality of that DNA. Other researchers have proposed incorporating multiple probes to assess gender, degradation and inhibition (Hudlow, Chong et al. 2008). At least one researcher has evaluated the incorporation of multiple sized amplicons targeting both nuclear and mitochondrial DNA (Niederstatter, Kochl et al. 2007). Although, as noted in the RNA section above, some research has been conducted investigating the application of RT-qPCR combined with Reverse Transcriptase RT-PCR of RNA to assess time since death, using RT-qPCR to compare of the rate of decomposition between tissues (Trindade-Filhoa, Mendesa et al. 2008), and determining insect age to assess time since death (Ames, Turnera et al. 2006), little or no work has been performed applying it to the assessment of time since death.

“Fragment Size Analysis”

Unlike both competitive PCR and RT-qPCR, both processes initially implemented as quantitative PCR techniques then adapted by investigators to assess degradation, the process of evaluating the size of amplified fragments to assess DNA fragmentation, hereafter referred to as Fragment Size Analysis (FSA) PCR, was specifically developed to allow the estimation of DNA “quality” and provides little information about the absolute DNA quantity. In this instance quality can be defined as a combination of the fragmentation level of the DNA as well as it's availability for amplification. As noted above, DNA that is fragmented to the point where little or no intact fragments span the desired amplicon will produce little or no amplified product. Additionally, samples with intact DNA but inhibited by contaminants that bind the DNA or co-purify with the DNA may also fail to amplify. Unlike quantitative PCR methods which typically utilize smaller amplicons to provide estimates of the absolute quantity of DNA; but can underestimate the quantity of DNA in the larger amplicon ranges; FSA PCR focuses on assessing larger amplicons. Additionally, unlike RT-qPCR an internal positive amplification control is used with each sample and where the amplicon length is ideally restricted to 50 - 150bp (Ambion 2006), or competitive PCR where the competitor DNA must be pre-amplified, purified, quantified then added to the amplificaiton (Zimmermann and Mannhalter 1996); FSA PCR is less complicated in that it does not require an internal standard/competitor and the fragment length is limited only by primer design, enzyme efficiency and detection method.

Several investigators have developed FSA PCR assays specifically do assess DNA quality in forensic samples. One proposed assay used fluorescently labeled amplicons in the mid-range of STR analysis (164bp) and mitochondrial sequencing (260bp) assayed on a ABI Prizim® 310, to predict the likelihood that a forensic sample (bone, formalin fixed tissue and casework) would produce a full STR profile (von Wurmb-Schwark, Schwark et al. 2004). Another researcher used the amplification of fragments of the beta-actin gene ranging from 150-750bp to assess the quality of DNA in various layers of bone ranging in age from 1-200 years (Kaiser, Bachmeier et al. 2008). Most recently, researchers in Australia used FSA PCR to examine effect of time and temperature on porcine skeletal muscle (Larkin, Iaschi et al. 2009).

Research Objectives

Forensic scientists that perform DNA analysis are not only interested in the application of molecular biology techniques to the area of forensic DNA analysis; but, like all forensic scientists, are also interested applying existing technology in their field to new areas of forensics. It has been well established that the degradation of DNA impacts its ability to be typed using standard genotyping methods (Clayton, Whitaker et al. 1995; Dupuy and Olaisen 1997). Further, studies using the controlled enzymatic degradation of purified DNA have shown that the capacity of a genotyping method to produce a profile is directly related to its minimum required DNA fragment length and the level of degradation, thus it varies from procedure to procedure and specimen to specimen. Such studies have been invaluable in establishing the limitations of a genotyping method related to the level of degradation exhibited by the DNA specimen under controlled conditions. Additional studies, examining the effects of various environmental insults have been performed on degraded tissues in order to shed light on the strengths and limitations of the genotyping method examined (Kimpton, Oldroyd et al. 1996; Sparkes, Kimpton et al. 1996; van Oorschot, Gutowski et al. 1996). Although instrumental in developing and understanding new genotyping methods, these studies have been limited to “naked” DNA, tissue specimens degraded in uncontrolled conditions or stains on cloth. While such samples simulate certain types of DNA evidence found at crime scenes, as with the enzymatic degradation studies, these samples typically are not degraded under controlled conditions, and thus are of limited value as models in studying post mortem interval. Finally, unlike anthropological studies designed to assess degradation in specimens under strictly controlled environmental conditions to determine post mortem interval (Di Nunno, Costantinides et al. 1998), most DNA validation studies that examine degraded DNA have not attempted to correlate typing data with post mortem interval.

This study was designed to examine the effect of temperature and time on the degradation of DNA within soft tissue, and to determine if those effects could be used to estimate post mortem interval. The domestic pig, Sus scrofa scrofa (referred to by some taxonomists as S. domestica) was selected as the human tissue analog for this study. Domestic pigs have been widely used in forensic anthropology studies. Although investigators have questioned the use of domestic pigs as a human tissue analog (Jonuks and Konsa 2007) others have found pigs to be reliable models for human corpses (Schoenly and Hall 2002; Hughes and White 2009; Stokes, Forbes et al. 2009). Like humans, pigs are omnivorous, possess a similar digestion system, similar intestinal flora and similar distributions of body hair and skin tissue (Micozzi 1991; Dillon and Anderson 1996). In addition to the convenience of being able to regularly obtain samples, the use of pigs and pig tissue prepared for food eliminates regulatory issues associated with the Animal Welfare Act of 1966 and the Human Tissue Act of 1983.

To accomplish the objectives of this project the following aspects were addressed:

1. Suitable loci/markers were identified.

2. A PCR-based assay that allows the assessment of the fragmentation level of DNA was developed.

3. A method of controlling or eliminating microbial putrefaction was established.

4. Skeletal muscle was degraded and analyzed under the controlled conditions.

5. The relevant data was analyzed.

Oligonucleotide primers were designed to allow the PCR amplification of varying sized fragments of nuclear loci, mitochondrial loci and RAPD markers. An FSA PCR based assay was developed using DNase degraded DNA. Tissue samples, handled a manner designed to decrease or eliminate the microbial putrefaction, were degraded under laboratory controlled conditions and assessed with the FSA PCR assay. The amplified fragments, generated from sample exposed to varying conditions, were analyzed using gel electrophoresis. Differences in the intensity of the amplified products were compared and assessed.

The application of FSA-PCR testing on samples degraded in a controlled fashion should provide a valuable technique for assessing time since death based on DNA degradation. However, even if the method is unable to provide definitive estimates of the postmortem interval, it should provide additional information regarding the rate of DNA degradation related to autolysis as well as providing another method to assess the effects of varying conditions on DNA degradation.

Chapter II
Materials and Methods

Tissue selection

DNA and tissue used for all laboratory studies were prepared from muscle tissue (tongue). Tongues were collected from a local abattoir at or near (within 1 hour) of the time of slaughter and stored on dry ice for not more than 1 hour before hard freezing at -80°C. Tissue used for all field samples included skin and blood collected from an intact animal. An animal was collected from a local abattoir at the time it was sacrificed using a penetrating captive bolt pistol. The head wound was sealed with silicone caulk and the animal transferred to a partially shaded location and placed in a wire mesh cage within two hours of death.

Field location

Tissue subsamples were placed at 33.130694° N, 97.160127° W 689 ft elevation with temperatures collected at 33.206° N, 97.199° W 676 ft elevation Intact animal field samples were placed and collected at 36.25877° N, 86.446524° W 479 ft elevation with temperatures collected at 36.125674°N86.678009°W 554 ft. Coordinates and elevations were plotted using Google maps and www.heywhatsthat.com.

Tissue Sampling

Tissue samples used for laboratory assessments were collected from frozen pig tongue using a 0.75 cm diameter brass cork boring tool. Previous studies indicate that freezing the tissue prior to use does not affect decomposition (Stokes, Forbes et al. 2009). For sampling, the frozen tongue was removed from the freezer and placed on a surface pre-sterilized with a 1:10 dilution of regular strength (5.25%) household bleach (sodium hypochlorite). The sterilized cork boring tool was used to remove a 2-3 inch plug of tissue. A 2-3 mm circular section was removed from end of the plug corresponding to the outer dermis of the tongue. The still frozen plug was placed in a pre-sterilized 50 ml screw-cap conical tube and placed in the -80°C freezer until needed. Sub samples of the tissue plug were prepared using a clean plastic weigh boat, clean razor blades and heat sterilized forceps. A 1-3mm circular section (approximately 0.1g) of tissue was removed from the plug. Once a section was removed from the plug it was placed into a pre-sterilized 1.5 ml microcentrifuge tube or a pre-sterilized 15 ml screw-cap conical tube.

Field samples collected from the intact animal were collected using three different procedures. Reference blood samples were collected from the head wound produced during the animal sacrifice using 4 - 4 inch sterile Dacron® applicators. The blood was allowed to completely dry at room temperature and were stored 15 ml screw-cap conical tubes at -20°C until needed. During the initial stages of decomposition (days 1-10) tissue samples were collected from the intact field specimen using a clean 0.75 cm diameter brass cork boring tool. An approximately 1-3 cm long plug was removed from muscle tissue by driving the boring tool through the dermis into underlying muscle. Once the plug was removed the hole was plugged with silicone caulk. Subsequent samples were collected approximatly 0. 5 cm away from previous sample sites. The tissue plug was placed into a pre-sterilized 50 ml conical tube with approximately 10 ml of isoproponal, transferred at room temperature to an intermediate storage freezer (-7°C) for not more than 24 hours and transferred to a long-term storage freezer (-20°C) until sub-sampled for analysis. Sub samples of the tissue plug were prepared as noted above. During the second phase of decomposition (~days 11-25) drying of the dermis, breakdown in muscle connective tissue and insect activity made penetration of the skin and successful muscle tissue collection difficult using the cork boring tool. An alternative method using a clean forceps and scissors was used. The dermis was lifted away from the underlying tissue and scissors were used to excise a circular portion of the dermis and any adhering tissue. Once the section was removed the hole was plugged with silicone caulk. Once sections were collected the hole was plugged with silicone caulk. During the third phase of decomposition (~days 26-35) tissue breakdown and insect activity had advanced to the point that silicone plugs and “patches” failed to adhere to the surrounding tissue. Samples collected during the third phase of decomposition were collected using the same procedure as phase two samples. However, no attempt was made to close tissue collection sites. The tissue sections were stored as noted above.

There were several issues noted with collection. Narrow rope or wire attached to the limbs at the outset of the experiment would make re-positioning of the specimen easier when shifting due to bloat occurs. On-site high/low temperature data was lost for 18 days. The site was mowed 15 days after specimen placement.

Genomic DNA isolation

There are multiple procedures available for the extraction of high quality DNA from tissue and dried blood samples. For the purposes of this study two different isolation procedures were evaluated. Isolation procedure 1, a DNA extraction technique commonly used in forensic applications, was used early in the research and was found to produce suitable yields of genomic DNA but unsuitable quality (highly degraded). Isolation procedure 2, while more complicated and time consuming, consistently produced DNA of the quantity and quality necessary to complete these studies.

Isolation procedure 1

Tissue sub-samples were prepared from tissue plugs or tissue sections using clean scissors or a disposable scalpel to collect a small portion of the sample (approximately 50 mg) from the tissue bore or cut section. While still frozen the sub-sample was then sliced into wafer thin slices and placed a microcentrifuge tube.

A 500µl aliquot of stain extraction buffer (10mM TrisHCl, 100mM NaCl, 10mM EDTA, 2% SDS) combined with 25µl Proteinase K (20mg/ml) and 50µl 0.39M DTT was added to each sample tube. The sample was vortexed to mix and centrifuged at approximately 14,000 rpm (top speed) for 5 - 10 seconds to force the solution and tissue sections to the bottom of the sample tube. Samples were incubated in a 56°C water bath for 24 hours*. Proceed to organic extraction.

Isolation procedure 2

Tissue sub-samples from plugs or sections were removed using clean razor blades and forceps sterilized with bleach prepared as noted above. A section of tissue or plug (approximately 100mg-500mg) was dropped into a mortar cooled to -196°C with liquid nitrogen and maintained at -79°C in a bed of crushed dry ice. Liquid nitrogen was poured into the mortar over the tissue being careful not to wash the tissue out into the dry ice bed. The tissue, floating in the liquid nitrogen, was ground into powder using a pre-cooled pestle.

The tissue powder was scraped/poured into a 15ml conical tube and mixed with 2ml of tissue digest buffer (100mM NaCl, 10mM Tris-HCL pH8, 25mM EDTA pH8, 0.5% SDS tissue digest buffer). Proteinase K was added to a final concentration of 0.1mg/ml (Ausubel 1987). The samples were digested overnight at 56°C in a shaking waterbath. The samples were gently inverted to mix several times in the first few hours. Samples with more than 200mg tissue were supplemented with an additional aliquot of Proteinase K after six hours of digestion.

DNA Purification/Concentration

Organic Purification:

Samples were removed from the water bath and centrifuged for 3 minutes at 1000 rpm to force any condensation to the bottom of the tube. An equal volume of phenol:chloroform:isoamyl alcohol was added to the tube. The mixture was gently vortexed to achieve a milky emulsion and centrifuged for 5-15 minutes at 1,000 rpm. The upper aqueous layer was transferred to a 15ml siliconized chlorex tube with a new transfer pipette.

A 1/10 volume of a 3M sodium acetate (NaOAC) solution was added to the aqueous phase and swirled to mix. Two volumes of cold 100% ethanol were added to the tube. The tube was sealed with parafilm and mixed by gently inverting the tube. The parafilm was pierced with a pipette tip and the tube was placed in the -80°C freezer for a minimum of one hour. The tube was removed from the freezer and centrifuged for 10 - 20 minutes at 7000 x g. The ethanol was immediately decanted taking care not to dislodge the pellet. The pellet was washed by the addition of 1 ml ice-cold 70% ethanol, inverting once, and centrifuging for 10 - 20 minutes at 7000 x g. The ethanol was immediately decanted, again taking care not to dislodge the pellet. The inverted tube was blotted on a kimwipe and placed upright in a tube rack. The rack was placed in a vacuum concentrator for 10 minutes or until the pellet was dry. The tube was removed from the vacuum concentrator and the pellet suspended in 100 - 200ul of sterile deionized water. The DNA solution was further processed with either Microcon microfiltration or Qiagen Qia-amp column purification.

Microcon 100® Microfiltration:

A variety organic and inorganic molecules can co-purify with DNA and inhibit PCR amplification (Peist, Honsel et al. 2001; Bessetti 2007). In order to reduce potential inhibitors extracted samples were filtered using microfiltration. A Microcon 100® microfiltration device was assembled and labeled with the sample number. The Microcon 100® membrane was pre-wetted by the addition of 100ul of sterile dionized water to the top of the membrane in the membrane unit. The water was spun through the membrane by a 2-3 minute centrifugation at 3500 rpm in a microcentrifuge. The extract was transferred to the Microcon 100® membrane and centrifuged in a microcentrifuge at 3500 rpm for 10 minutes or until the liquid has filtered though the membrane. The membrane unit was removed from the Microcon 100® assembly and the filtrate discarded. The Microcon 100® was reassembled and 200ul of sterile deionized water was added to the membrane unit as a wash step. The water was centrifuged in a microcentrifuge at 3500 rpm for 10 minutes or until the liquid has filtered though the membrane. The wash step was repeated at least on time. One hundred microliters of water was added to the membrane unit and the membrane unit was removed from the assembly and inverted into a retenate capture tube labeled with the sample number. The retentate assembly was centrifuged in a microcentrifuge at 3500 rpm for 5 minutes. The membrane unit was discarded. The filtered DNA was stored at -20°C until used.

Qiagen QIAamp Mini-Column Purification:

In order to improve the DNA recovery and remove potential PCR inhibitors, DNA extracts were re-purified using the QIAamp Mini® columns. After the drying step of the organic extraction protocol the DNA pellet was suspended in 160μl of QIAamp ATL digest buffer was added to each sample. A 40μl aliquot of RNase A (10mg/ml) was added to the DNA solution. The sample was mixed by vortexing for 15 seconds and incubated at room temperature for 2 minutes. The sample was centrifuged at low speed to remove liquid from the inside of the lid. After centrifugation 200μl AL buffer was added to the tube. The sample was mixed by vortexing for 15 seconds and incubated at 70°C for 10 min. The sample was centrifuged at low speed to remove liquid from the inside of the lid. After centrifugation 200μl of ethanol (96-100%) was added to the sample and mixed by vortexing for 15 seconds. The sample was centrifuged to remove liquid from the inside of the lid. The mixture, and any precipitate that may have formed, was applied to the QIAamp Mini® spin column. The column was centrifuged at 6000 x g (approximately 8000 rpm) for 1 minute. The QIAamp Mini® spin column was transferred to a clean 2 ml collection tube and initial filtrate tube and binding solution was discarded. After column transfer 500μl of QIAamp buffer AW1 wash buffer was added to the column. The column was centrifuged at 6000 x g (8000 rpm) for 1 minute. The QIAamp Mini® spin column was transferred to a clean 2 ml collection tube and the initial wash filtrate tube and wash solution was discarded. After column transfer, 500μl of QIAamp AW2 wash buffer was added to the column. The column was centrifuged at 20,000 x g (approximately 14,000 rpm) for 3 minutes. The QIAamp Mini® spin column was transferred to a clean 1.5 - 2ml microcentrifuge tube and the second filtrate tube and wash solution was discarded. The DNA was eluted from the column by adding 200μl of distilled water, incubating at room temperature for 5 minutes then centrifugation at 6000 x g (approximately 8000 rpm) for 1 minute. The elution step was repeated with an additional 200ul of distilled water. Eluted DNA was stored at -20°C until used.

Quantitative Characterization of DNA Samples

The quantification (concentration determination) of nucleic acids is paramount for optimizing the dsDNA and/or ssDNA to enzyme ratio utilized in many molecular procedures. Several methods were utilized to measure the nucleic acid concentration.

UV Absorbance A220 - A320

The most commonly used technique for assessing the concentration of nucleic acids is measurement of UV absorbance. Advantages to using UV absorbance for determining concentration are that it is fast, accurate and does not require expensive reagemts. However, absorbance values are adversely affected by the presence of proteins, phenol and other contaminants. Additionally, the process is not it is not very sensitive at very low levels and is unable to differentiate between ssDNA, dsDNA and RNA. DNA and RNA exhibit maximum absorbance at ~260 nm. Due to tryptophan residues, proteins exhibit a maximum absorbance at ~280 nm. Phenol exhibits a maximum absorbance at ~270 nm. The ratio of absorbance at 260 nm: 280 nm is often used as a means to assess DNA purity.

When used to assess DNA an A260 reading of 1 corresponds to a 50 ug/ml dsDNA or 33 ug/ml for ssDNA.

When oligonucleotides were received from the manufacturer (generously provided by Bio-Synthesis Inc.) they were diluted and assayed with a BioRad spectrophotometer. A cursory analysis to determine yield and quality of the newly acquired oligonucleotide was routinely performed. First the oligonucleotide was reconstituted from its desiccated form in ddH20 (330 µl for 5 OD units), and then was diluted 1:100 by placing 10 µl oligonucleotide solution into 990 µl ddH20 for spectrophotometer analysis. The diluted sample was then placed into a 1 ml quartz cuvette for a wavelength scan from 220 nm to 320 nm. An absorbance reading of 1.0 in the range of 256 nm to 260 nm was assumed to represent approximately 33 µg/ml of single-stranded (oligonucleotide) DNA.

Experimental samples they were diluted and assayed with a BioRad spectrophotometer. A cursory analysis to determine yield and quality of the newly acquired oligonucleotide was routinely performed. First the sample was diluted to either 1:100 or 1:10 by placing 10 µl or 100 µl of sample into 990 µl or 900 µl ddH20 for spectrophotometer analysis. The diluted sample was then placed into a 1 ml quartz cuvette for a wavelength scan from 220 nm to 320 nm. An absorbance reading of 1.0 in the range of 256 nm to 260 nm was assumed to represent approximately 50 µg/ml of duplex DNA. Phenol or protein contaminants are known to shift the absorbance maximum to the right (270 nm - 280 nm). Samples exhibiting a contamination shift were re-purified and re-quantified.

Fluorescent DNA Quantitation

Early in the project experimental dsDNA samples were quantified using a PicoGreen® ([2-[N-bis-(3-dimethylaminopropyl)-amino]-4-[2,3-dihydro-3-methyl-(benzo-1,3-thiazol-2-yl)-methylidene]-1-phenyl-quinolinium]+) based quantification assay (Trudeau 2004). PicoGreen® is a fluorescent nucleic acid stain for quantifying double-stranded DNA (dsDNA) in solution. The structure of PicoGreen® (Ygonaar 2006)is such that once bound to dsDNA, it becomes intensely fluorescent exhibiting a 1000-fold increase of signal. The increase of signal allows samples ranging from 25 pg/ml - 1000 ng/ml to be quantitated using a standard fluorescence microplate reader (Singer, Jones et al. 1997).

Fluorescent quantification calibrators were prepared from a 50 ng/ul dsDNA stock solution and sterile distilled water in the following concentrations:

Cal-0.0 ng/ul

Cal-10.0 ng/ul

Cal-5.0 ng/ul

Cal-2.5 ng/ul

Cal-1.25 ng/ul

Cal-0.625 ng/ul

A 1:500 working stock of PicoGreen® was prepared using sterile distilled water. An optically clear scanning plate was prepared with 4ul of calibrator or experimental dsDNA. The scanning plate was incubated for five minutes at room temperature, protected from light. After incubation was complete, the samples were exposed to fluorescent light and the Relative Florescence Units (RFU) was measured using a using a fluorescence microplate reader. RFU measurements were obtained using a BMG Labtech FLUOstar fluorescent plate reader with excitation and emission wavelengths of ~480 nm and ~520 nm respectively. The plate was continuously agitated for 300s and the fluorescence of each well assayed five times with the reported value representing an average RFU value. The fluorescence value of the reagent blank was subtracted from the calibrator and sample values. The corrected calibrator data was used to generate the standard curve of RFU versus dsDNA concentration. Relative fluorescence measurements of samples with unknown DNA concentrations were compared to the standard curve and assigned concentration values using regression analysis. The calibrators were verified by assessing the r2. Calibrator sets with r2 values less than 0.98 were repeated.

Quantitative/Qualitative Characterization of DNA Samples

Agarose gel separation was used to perform quantitative and qualitative DNA analysis of some dsDNA samples. After separation nucleic acids were stained using ethidium bromide (EtBr), a dsDNA intercalating agent. After staining, gels were evaluated by the use of a UV transilluminator, a hand-held UV light source or by scanning with the BioRad Molecular Imager FX. A visual comparison of the experimental samples to calibrator samples and/or ladders allowed for the estimation of fragment size, quality and DNA concentration based on the shape, size and location of the stained bands/material on treated gels.

Sizing Ladders:

Three primary DNA sizing ladders were used for this project. The Hinf I-digest of pBR322 was used during the initial assay development stages in both agarose and acrylamide gels. This digest includes ten discrete fragments including 1632bp, 517bp, 504bp, 396bp, 344bp, 298bp, 221bp, 220bp, 154bp and 75bp fragments. The Fermentas ZipRuler™ Express DNA ladder Set was used in some agarose and all acrylamide gels during the analysis of all samples degraded under laboratory controlled conditions. The ZipRuler™ Express DNA Ladder Set consists of two ladder sets composed of chromatography purified DNA fragments. The first set, used as the primary ladder in this study, is ZipRuler™ Express DNA Ladder 1 which consists of 100bp, 300bp, 500bp 850bp 1200bp, 2000bp, 3000pb, 5000bp, and 10000bp fragments. The second set is ZipRuler™ Express DNA Ladder 2 consisting of 200bp, 400bp, 700bp 1000bp 1500bp, 2500bp, 4000pb, 7000bp, and 20000bp fragments. The Invitrogen trackit™ 100 bp ladder is prepared from a plasmid containing repeats of a 100bp DNA fragment. The Invitrogen's Trackit™ 100bp ladder consists of 100bp, 200bp, 300bp, 400bp, 500bp, 600bp, 700bp, 800bp, 900bp, 100bp, 1100bp, 1200bp, 1300bp, 1400bp, 1500bp and 2072bp fragments.

Preparation of Ladders, Calibrators and Experimental Samples:

Prior to mixing with loading dye, ladders, calibrators and experimental samples were gently vortexed and centrifuged for 5 seconds at full speed to mix and bring the contents to the bottom of the tube. Depending on the final volume prepared (10µl-20µl), 2µl-4µl of 5X loading buffer (10 mM Tris-HCl; 1 mM EDTA, pH 7.5; 0.005% bromophenol blue; and 0.005% xylene cyanol FF) was transferred to the bottom of a labeled microcentrifuge tube PCR amplification plate well or PCR amplification tube. Transfer 2µl-4µl of experimental sample, 2µl-4µl calibrator DNA, or 2µl-5µl Ladder to the loading buffer in the corresponding tube/well. Sample, calibrator and ladder volumes were adjusted to a final volume of 10µl-20µl by the addition of 4µl-14µl sterile distilled water to the bottom of the corresponding tube/well.

Preparation of Agarose Gel Solution:

Agarose gels were prepared by mixing agarose in a weight to volume ratio of 1.2g:100ml LE Agarose to 1XTris Borate EDTA (TBE) buffer. Agarose gel stock was prepared in batches by mixing 4.8g SeaChem LE Agarose with 40ml 10XTBE or 80ml 5XTBE in a 500ml bottle or flask. Adjust the final volume to 400ml using distilled water (makes 4 100ml gels). Swirl gently to mix the contents.

Autoclave Melting Method:

The bottom of chamber of the autoclave was filled with deionized water until the level was just below the bottom ledge of the door. The bottle(s)/flask(s) of agarose were placed in the autoclave chamber. The door was closed, latched and the chamber exhausted on the slow setting. The agarose was autoclaved for 25 minutes. The agarose was allowed to cool at room temperature until the boiling stopped. The agarose was gently swirled to mix. The agarose was either used immediately to pour gels or cooled to room temperature and stored at 4°C until used.

Microwave Melting Method:

The bottle(s)/flask(s) of agarose were placed in microwave and run on high for 6-8 minutes to boil. The agarose was allowed to cool at room temperature until the boiling stopped. The agarose was gently swirled to mix. If the agarose was not completely melted the solution was heated for one minute intervals until the agarose was melted. The agarose was cooled at room temperature until the boiling stopped. The agarose was either used immediately to pour gels or cooled to room temperature and stored at 4°C until used.

Casting Gel:

The agarose gel form was created using tape to seal the ends of an acrylic tray. One hundred grams of agarose gel stock was placed into a 250ml flask and heated in a microwave until the agarose was completely melted. A 15µl aliquot of 1% ethidium Ethidium_bromide.pngbromide(Ygonaar 2006) was added to the melted agarose which was swirled to mix. The melted agarose solution was poured into the gel tray. A comb was placed at the top of the gel if needed in the middle position and the agarose was allowed to solidify for 30 minutes. Once the gel solidified, the tape was removed from the edges of the gel tray and the gel placed into the electrophoresis tank with the comb at the negative (black electrode) end of the gel. The gel tank was filled to just above the level of the gel surface with 1X TBE buffer. The comb(s) were removed taking care not to tear the wells.

Gel Loading:

The ladders, calibrators and/or samples, prepared as noted above, were individually loaded into separate gel wells.

Electrophoresis:

The gel tank lid was placed on the tank and the wire leads, red to (+) positive and black to (-) negative, were connected to the power supply. The current was set from 100 - 200 volts and the gel was run from 10 - 30 minutes.

Interpretation:

Once electrophoresis was complete the gel was removed from the gel box and placed on a UV transilluminator. The gel was documented by photograph Polaroid® 667 black and white film (1 second exposure at f4.5 with an orange filter) or digital documentation.

Invitrogen EGel

E-Gel® pre-cast agarose gels are self-contained agarose gels that include electrodes packaged inside a dry, disposable, UV-transparent cassette. The E-Gel® agarose gels run in a specially designed device that is a base and power supply combined into one device (two bases are available for running E-Gels, the new iBase™ system and the original, economical E-Gel® Powerbase™).

Pre-electrophoresis:

The E-Gel® was removed from the package and inserted (with the comb in place) into the E-Gel® Powerbase™ right edge first. Proper placement required that the Invitrogen logo be located at the bottom of the E-Gel® Powerbase™. The E-Gel® was pressed firmly at the top and bottom to seat the gel in the E-Gel® Powerbase™. The E-Gel® PowerBase™ v.4 was pluged into an electrical outlet using the adaptor plug on the base. A steady, red light on the base was activated when the gel was correctly inserted. The gel was pre-run (with the comb in place) by pressing and holding either the 15 min or 30 min button until the red light turned to a flashing-green light indicating the start of a 2-minute pre-run. At the end of the pre-run, the current automatically shut off. The combs were removed from the E-Gel® cassette.

Gel Loading:

The ladders, calibrators and/or samples, prepared as noted above, were individually loaded into separate gel wells.

Electrophoresis:

The 30-minute or 15 minute button on the E-Gel® PowerBase™ was pressed to begin electrophoresis. At the end of the run the current automatically shut off and the power base signaled the end of the run with a flashing red light and rapid beeping. Either button was pressed to stop the beeping. The gel cassette was removed and analyzed.

Interpretation:

Remove the gel from the gel box and place on the UV transilluminator. Photograph the gel with Polaroid 667 Black and White film for 1 second at f4.5 with an orange filter. If the gel image is too dark, you may need to increase the time to 2 seconds. If the gel image is too bright, you may need to decrease the time. Develop the picture for approximately 45 seconds to ensure that you have a quality photograph.

Preparation of degraded DNA:

The second step of degradation was achieved by DNAse I treatment in a Mn2þ-buffered (50 mM Tris-HCl, pH 7.6; 10 mM MnCl2) solution. Pancreatic DNAse I normally introduces single-strand nicks into doublestranded DNA. In the presence of Mn2þ, DNAse I cleaves both strands of DNA at approximately the same site [8]to yield fragments of DNA that are blunt-ended or that have protruding termini only one or two nucleotides in length. Sonicated DNA was digested with DNAse I at 15 ng/ml and after defined time periods aliquots of 15 ml were taken and the reaction was stopped by adding 7.5 ml of 50mM EDTA solution. The recovery of DNA was monitored by electrophoresis in an ethidium bromide-stained 1.5% agarose gel and DNA concentration was estimated by fluorometry. (Bender, Farfan et al. 2004)

Degraded DNA samples were prepared by treating previously quantified high molecular weight dsDNA control samples with pancreatic DNase I. Control samples were treated with DNase I in a Mg2+-buffered solution to produce dsDNA fragments of varying sizes. Pancreatic DNase I introduces random single-strand nicks into dsDNA. Digestion was allowed to proceed for varying lengths of time from 0 seconds to 120 minutes. A standard DNase I digestion was set up as follows:

1µl 100mM Tris-HCl, pH 8

1µl 10mM MgCl2; 1mM CaCl2

5µl DNA (0.1µg/µl)

2µl Sterile Distilled Deionized Water

1µl Pancreatic DNase I (2ng/ µl)

10µl Total Reaction Volume

DNase I was inactivated by the addition of 10µl stop solution and heating (95°C for 10 minutes). Stop solution was prepared as follows:

2µl EDTA 100mM

8µl Sterile Distilled Deionized Water

Digested DNA was stored at -20°C until used.

RAPD analysis

Nine random 10-mer primers used for RAPD analysis (Ramser, Weising et al. 1997) were ordered from Applied Biosystems.

Primer 1* 5'-GAACGGACTC-3'

Primer 2* 5'-AAAGCTGCGG-3'

Primer 3* 5'-TGTCATCCCC-3'

Primer 4* 5'-CACACTCCAG-3'

Primer 5* 5'-GTTGCCAGCC-3'

Primer 6* 5'-GTGCCTAACC-3'

Primer 7* 5'-TCACGTCCAC-3'

Primer 8* 5'-CTCTCCGCCA-3'

Primer 9* 5'-GGATGAGACC-3'

* = 5' labeled with 6-FAM

PCR amplifications were performed in the following fashion. A new set of thin-wall (PCR) reaction tubes was set up with each containing the following:

10µl DNA (3ng/µl)

2.5µl Promega Gold ST*R 10x Buffer™ (w/dNTP Mix)

2.5µl RAPD primer 1-9 (10µM)

0.2µl Taq polymerase (5 U/µl)

9.8µl ddH2O

25µl Total volume

The parameters for the RAPD thermal cycling reaction were as follows:

94°C for 11 minutes

3 cycles of 94°C for 30 seconds

34 °C for 30 seconds

72°C for 90 seconds

35 cycles of 94°C for 15 seconds

34 °C for 30 seconds

72°C for 90 seconds

72°C for 7 minutes

4°C hold

The resulting PCR amplification products were analyzed by gel electrophoresis.

Directed PCR amplification

Primers for the directed amplification of Group Component (Jorgensen, Hobolth et al. 2005) were ordered from Operon.

Primer GC A* 5'-aggacttgccagcagaaaaa-3'

Primer GC B1 5'-tctggaagctcaggtcttgg-3'

Primer GC B2 5'-ccaaattcgcagtaggcact-3'

Primer GC B3 5'-ccttggggtctttcctgaat-3'

Primer GC B4 5'-agcaggcttctgtcaaggac-3'

Primer GC B5 5'-tgctacagcaagcaggaaaa-3'

* = 5' labeled with fluorescein (FL)

PCR amplifications were performed in the following fashion. A new set of thin-wall (PCR) reaction tubes was set up with each containing the following:

1µl DNA (3ng/µl)

1µl Promega Gold ST*R 10x Buffer™ (w/dNTP Mix)

0.1-0.4µl Directed Primer Left (10µM)

0.1-0.4µl Directed Primer Right (10µM)

0.2µl Taq polymerase (5 U/µl)

7-7.6µl ddH2O (volume depends on primer quantity)

10µl Total volume

The parameters for the directed PCR thermal cycling were as follows:

94°C for 4 minutes

29 cycles of 94°C for 30 seconds

64°C for 45 seconds

72°C for 1 minute

72°C for 7 minutes

4°C hold

The resulting PCR amplification products were analyzed by gel electrophoresis.

Vertical Non-Denaturing Polyacrylamide Gel Electrophoresis with DNA degradation has been identified by a loss of high molecular weight DNA on agarose gels, by a decrease in the concentration of human DNA to total DNA using agarose gels combined with slot-blot analysis, by the drop out of alleles at larger loci with RFLP, Variable Number of Tandem Repeats (VNTR) and Short Tandem Repeats (STRs), and by the shift in Ct curves using real-time PCR (Frank and Llewellyn 1999; Fujita, Kubo et al. 2004; von Wurmb-Schwark, Schwark et al. 2004).

Separation of dsDNA created by PCR amplification was routinely performed using non-denaturing polyacrylamide gel electrophoresis. N-PAGE procedures were optimized to allow differentiation of amplicons ranging from 50bp - 1500bp. The procedures to assemble the polyacryamide gel cassette is outlined below.

Different gel cassette sizes and acrylamide concentrations can be utilized for polyacrylamide gels, but only one was used in these experiments. The 20 x 20 cm, cassette was used to verify the presence of PCR products. Two glass plates, one 20 x 20 cm and the other 20 x 22 cm, were soaked for 30 minutes in 1M NaOH to strip contaminants from the plates. After stripping the plates were washed with alconox detergent and distilled water. Once dry, both plates were coated on one side with a thin layer of 5% dichloro-dimethylsilane dissolved in heptane. The plates were allowed to dry and were then wiped with Kimwipes® before completing the cassette assembly. The silinating was performed in a vented hood to prevent inhalation of the potential carcinogen. Two spacers, 1.5 mm thick and 1.5 cm wide, were placed on either side of one of the plates. The other plate was then carefully placed on top of the spacers with clamps added to hold the plates together. The cassette assembly was placed on a flat taped across the bottom using yellow gel tape. Non-denaturing acrylamide gel solution was then prepared.

A ten milliliter volume of 40% acrylamide (29:1 acrylamide/bis-acrylamide) was added to a fifty milliliter conical tube. Ten milliliters of 5X non-denaturing buffer (500 mM Tris, pH set to 8.3 with solid boric acid, 10.0 mM EDTA, ethylenediaminetetra-acetate, disodium form) was mixed with the acrylamide/bis-acrylamide solution. Thirty milliliters of de-ionized water was used to bring the volume up to 50 ml, producing an 8% polyacrylamide solution. Ammonium persulfate, 0.06 g, was added to the solution and the mixture was filtered through a 1mm filter into a side-arm flask. A vacuum was applied (2-10 minutes) to de-gas the mixture. A polymerization catalyst, 8 µl TEMED (N,N,N',N'-tetramethylethylenediamine), was added and gently swirled before the solution was poured into the cassette using a 25ml glass pipette and powered vacuum pipetter. The comb was inserted after it was determined no bubbles were present in the assembly, which would inhibit polymerization. Clamps were used to clamp the comb, like a spacer, ensuring uniform gel thickness. The cassette was left in a horizontal position to prevent leakage and help promote uniform gel thickness. Polymerization was considered complete after 0.5 - 1 hour. The top of the cassette was wrapped with plastic wrap if it was to be allowed to sit overnight and the gel run the next day.

Prior to electrophoresis, the comb was removed and wells straightened and rinsed. The cassette was then placed into a vertical electrophoresis chamber and 1 liter of 1X non-denaturing buffer was equally dispensed into the upper and lower reservoirs. The gel was pre-electrophoresed for 10 minutes at 400 volts. One microliter of each PCR amplicon was mixed with two mciroliters of 5X loading buffer (15% ficoll, 0.1% bromophenol blue, 0.1% xylene cyanol FF, 50 mM EDTA) and brought up to ten microliters before loading in a washed well of the gel. The pBR322 HinfI-digested DNA was placed in two of the 18 wells as size standards. The amplified product was stacked and run into the gel by electrophoresis at 100 V for 15 minutes. The gel was then electrophoresed at a constant 400 V for 80 minutes.

DNA detection by SyberGreen/Gel Red Staining:

The buffer was drained from the reservoirs by inverting the unit over a sink. The gel cassette was then removed and disassembled to leave the gel on one plate, which was then submerged in a SyberGreen or Gel Red solution (10000:1 dilution) for 15 -30 minutes. The gel was then transferred using a 35 x 60 cm piece of flexible nylon window screening and transferred to a BioRad Molecular Imager FX.

DNA detection by SyberGreen/Gel Red Staining:

The buffer was drained from the reservoirs by inverting the unit over a sink. The

Temperature Tracking:

The daily temperature was monitored using temperature recordings obtained from a manual electronic device situated next to the carcasses, and published temperatures from the Meteorological Bureau at Geraldton Airport, a distance of some 30 km from the sampling site. The Geraldton Airport has the closest weather station to Drummond Cove. The daily maximum and minimum temperatures were recorded for 81 days during summer and 52 days during winter. The temperature was adjusted by subtracting the Drummond Cove average monthly temperature (maximum and minimum) from the average monthly temperature (maximum and minimum, respectively) taken by the meteorological bureau at Geraldton airport. This method was chosen as it is used in a number of forensic disciplines to calculate PMI.

Temperature data was collected from the nearest National Weather Service Station. Temperature date was accessed at the following websites:

http://www.rap.ucar.edu/weather/surface/

http://www.ncdc.noaa.gov/oa/climate/stationlocator.html

No temperature corrections were performed on temperature data used for calculations of Accumulated Degree Hours (ADH) or Accumulated Degree Days (ADD).

=

DH

The ADH were calculated by adding together the hourly values (Megyesi, Nawrocki et al. 2005; Gennard 2007).

The degree day value for a single day was calculated using the following formula:

Maximum Temperature + Minimum Temperature

-

Minimum Threshold

=

DD

2

The ADD were calculated by adding together the single day values (Larkin, Iaschi et al. 2009). For this study, a minimum thresholds of 0°C and 4°C were evaluated when calculating DH and DD (Micozzi 1991; Vass, Bass et al. 1992).

calculate this parameter providing mean values and confidence limits, including the precision and accuracy of calculation.

Statistical evaluation of the data involved least square analysis of linear regression. The scatter diagrams of time since death (dependant variable) were plotted against each of the three other (independent variables) viz. potassium, sodium and Sodium/potassium ratio. Then the data was linearized by using the model Y ¼ axb on a double logarithmic scale between log y, dependent variable and log x, independent variable. The model was: log y ¼ log a þ b logx

All logarithms were used to the base 10 where log a and b represents the intercept and scope of the regression line, respectively. These constants have been estimated from the data to predict the value of log y, i.e. the dependent variable. The actual predicted value of time since death may be obtained after taking the anti log.

Multiple regression analysis of the data was performed with every possible combination of the time of death and the NPN values in the tissues. (r = 0.673),

The design of the degradation experiments—comparing the three fragments with each other within a set of controlled conditions meant that the resultant quantitative data were examined in a relative, not absolute, manner. An advantage of this was that the amount of starting DNA added to a PCR reaction was not critical; the relative levels of the three loci within a sample (and the change in these ratios as samples degraded) were the important factor.


More from UK Essays

Get an Instant Quote - No registration required!