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Pyura DNA extraction

Pyura DNA extraction


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I'm am struggling with genomic DNA extraction from different samples of Pyura chilensis; the DNA is degraded as can be seen on the gel.

We've always used GeneJET Genomic DNA purification Kit (by ThermoFisher), and it works fine for samples from other species; so the kit isn't the problem.

We've always used tissue samples stored in 95% Ethanol (which are washed with water prior to DNA extraction), but the problem persists irrespective of whether the samples are one day old or a few years old. The storage method seems to work fine for gDNA extraction from other species.

Then, my guess is that the problem is with Pyura itself. So I have to figure out if either the extraction or the storage isn't appropriate for Pyura. I read that Pyura has a lot of copper in it. We planned to use a new buffer for storing it: salt Saturated DMSO, containing EDTA.

I would like to ask if any of you have ever worked with this organism (and if you have, then how did you manage to extract DNA from it). Do you have any suggestions on any DNA purification protocols?

1st lane is the ladder (1kb) and last 3 lanes with long blurred thing instead of bands are my DNA samples.


With reference to your gel photo, the genomic DNA (in the three Pyura sample lanes) does not look badly degraded. If the size marker is 1 Kb ladder, then the majority of the DNA is of high molecular size. During extraction it is common for high molecular weight DNA to become "sheared" to some degree (mechanically broken into smaller fragments). This can appear as the faint downward smear. The lanes may be somewhat over loaded as well. You can try to use more gentle extraction technique and see if it helps to your satisfaction. Any of those you mention will work.

Badly degraded genomic DNA would appear as a more intense "smear" near the lower portion of the 1 Kb size standard.


This is from a published work by Segovia et al. 2017:

Mantle tissue (0.2 g) from each individual was used to extract DNA using the DNeasy Blood® & Tissue Kit (QIAGEN®, USA) according to the manufacturer's instructions. Quantity/purity of DNA was measured in Nanodrop 2,000 (Thermo, USA).

They have done SNP identification from the genome. So I assume that their gDNA would have been of decent quality.


Pyura DNA extraction - Biology

Report of Systematic Zoology Lab Practicum, August, 2010

Cytochrome c oxidase subunit I partial sequence of a putative calanoid copepod obtained from Pyura vittata (Chordata: Ascidiacea: Pleurogona: Pyuridae)

Division of Biology, Department of Biological Sciences, School of Science, Hokkaido University, Sapporo 060-0810, Japan

Material and Methods
An ascidian was obtained subtidally at Oshoro Bay, Hokkaido, Japan, about 43°12&primeN, 140°51&primeE, on 31 May 2010 by Shin Hayase, photographed and identified by Hiroshi Kajihara as Pyura vittata based on Nishikawa (1992: 600, pl. 144-1) before fixed in 99% EtOH. DNA was extracted from the alimentary tract of the specimen, using the silica method (Boom et al. 1990) with some modifications. Extracted DNA was dissolved in 30 µl of deionized water and has been preserved at &ndash20°C. Remaining morphological voucher specimen has been deposited at the Hokkaido University Museum under the catalogue number ICHU22080146 (contact: Dr. Hiroshi Kajihara, [email protected]).
An about 600-bp fragment of mitochondrial cytochrome c oxidase subunit I gene (COI) was amplified by polymerase chain reaction (PCR) using LCO1490 (5&prime-GGTCAACAAATCATAAAGATATTGG-3&prime) and HCO2198 (5&prime-TAAACTTCAGGGTGACCAAAAAATCA-3&prime) (Folmer et al. 1994). A hot start PCR was performed by a thermal cycler, iCycler (Bio-Rad), in a 20-µl reaction volume containing 1 µl of template total DNA (approximately 10&ndash100 ng) and 19 µl of premix made with 632-µl deionized water, 80-µl Ex Taq Buffer (TaKara Bio), 64-µl dNTP (each 25 mM), 8-µl each primer (each 10 µM), and 0.1-µl TaKara Ex Taq (5 U/µl,TaKara Bio). Thermal cycling condition comprised an initial denaturation at 95°C for 30 sec 30 cycles of denaturation at 95°C for 30 sec, annealing at 45°C for 30 sec, and elongation at 72°C for 45°C and a final elongation at 72°C for 7 min.
The PCR product was purified with the silica method (Boom et al. 1990). Both strands were sequenced with a BigDye® Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems) following the manufacturer's protocol, using the same primer set as the initial PCR amplification. Sequencing was performed with ABI Prism 3730 DNA Analyzer (Applied Biosystems). Chromatogram and sequence data were operated with MEGA v4 software (Tamura et al. 2007).

Results and Discussion
A total of 606 bp of COI sequence was obtained. However, a nucleotide BLAST search of this sequence at National Center for Biotechnology Information indicated that the sequence probably came from a calanoid copepod, rather than the ascidian. It is concluded that a care must be paied to avoid alimentary tract for DNA extraction from ascidians.

Taxonomy
Phylum Chordata
Class Ascidiacea
Family Pyuridae Hartmeyer, 1908
Genus Pyura Molina, 1782
Pyura vittata (Stimpson, 1852)
(Fig. 1)



Fig. 1. Pyura vittata (Stimpson, 1852) (ICHU22080146), lateral view.

Boom, R., Sol., C. J. A., Salimans, M. M. M., Jansen, C. L., Wertheim-van Dillen, P. M. E., and van der Noordaa, J. 1990. Rapid and simple method for purification of nucleic acids. Journal of Clinical Microbiology28: 495&ndash503.

Folmer, O., Black, M., Hoeh, W., Lutz, R. and Vrijenhoek, R. 1994. DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Molecular Marine Biology and Biotechnology 3: 294&ndash299.

Nishikawa, T. 1999. Chordata. Pp. 573&ndash608. In: Nishimura, S. (Ed.) Guide to Seashore Animals of Japan with Color Pictures and Keys. Vol. II. Hoikusha, Osaka. xii+pls 73&ndash144+663 pp.

Tamura, K.,Dudley, J., Nei, M. and Kumar, S. 2007. MEGA4: Molecullar Evolutionary Genetics Analysis (MEGA) software version 4.0. Molecular Phylogenetics and Evolution 24: 1596&ndash1599.


Appendix
COI sequence from ICHU22080146 (identified as the ascidian Pyura vittata), though most probably representing a calanoid copepod.


This short activity helps students visualize one of the most important molecules on the planet, DNA. The activity can be done with simple materials found in most homes. We use bananas, but strawberries or other fruit soft enough to mush up can also be used. The activity is written for students at a middle school or higher level, but with more intense guidance, this activity is useful for students of any age.

Time Required : 45 minutes

  • One container of soap and salt should be more than enough for a large class to use, but we suggest having two of each on hand just in case.
  • This activity generally works better with small groups of students each working on their own banana extraction. This also makes it likely that at least one group will have very visible DNA.
  • Make sure to have extra zip bags and extra coffee filters on hand, in case any break.
  • If a coffee filter breaks and banana mush falls to the bottom of the glass, pour it back into the bag, secure a new filter, and pour the mush back in more slowly, letting it drain as you pour.

Extensions:

When combined with additional reading from Ask A Biologist, or additional short assignments, this DNA extraction activity can meet several learning standards.

  • The story “DNA ABCs” will help students understand the importance of DNA to life, as well as the chemical and physical structure of DNA.
  • The story page “Getting Genetics Straight” will help students differentiate between genes and chromosomes, and understand alleles.
  • For high school students, the story “Controlling Genes” will help them understand protein synthesis.
  • Students can try to figure out how much DNA is in an adult human body. You can either provide the following information to them, or have them search for it online:

      Cellular

      • The story “Building Blocks of Life” will help students understand the cell structure and function.
      • The story “Cells Living in Cells” will help students understand the different cell types that exist.
      1. Students will grasp that small molecules are tangible.
      2. Students will follow directions and understand that basic chemicals (salts and detergents) can be used to break down cells and cell parts and to make molecules stick to other molecules.
      3. EXTENSION: Students will gain a basic understanding of the structure of DNA.
      4. EXTENSION: Students will understand cells and that they broke apart the cell membrane and the nuclear membrane to reach the DNA.

      DNA Extraction Technique

      In this experiment, a goal is to extract the DNA from a fruit sample. Some knowledge of the scientific background behind DNA extraction is needed to do this.

      The DNA extraction process is a fairly simple biochemical procedure that can be divided into three major steps: breaking open the cell (lysis), destroying membranes within the cell, and precipitating the DNA out of the solution.

      The following sections describe how each step relates to the physical and biochemical properties of DNA.


      • DNA Extraction Buffer:1000 ml of deionized water, 50 ml of clear dish detergent, 1 teaspoon of salt
      • Strawberry (other fruits also work)
      • Ziploc bag, Coffee filters and funnels
      • Test tubes, beakers, or cups to collect filtrate
      • Ethanol or 91% isopropyl alcohol (chilled)

      1. Add a strawberry (or half) to a Ziploc storage bag.
      2. Add 10 ml of the DNA extraction buffer and mash the strawberry and buffer for about one minute.
      3. Use a funnel and coffee filters to filter the strawberry juice into a beaker.
      4. Transfer the filtrate to a test tube, you should only fill the test tube about half full and avoid transferring any foam.
      5. Slowly pour or drip cold alcohol over the top of the strawberry mixture. You want a single layer on top of the strawberry mixture.
      6. White strands will form in the ethanol layer, use a stirring rod or toothpick to spool the strands.


      Introduction

      With thousands of described species, ascidians, or sea squirts (phylum: Chordata, subphylum: Tunicata, class: Ascidiacea) form a unique group of sessile marine non-vertebrate chordates (Shenkar and Swalla 2011 Shenkar et al. 2012). Because of their key systematic position as a vertebrate sister-clade (Delsuc et al. 2006 Singh et al. 2009), ascidians have a pivotal role in evolutionary developmental studies and have become important animal models in comparative genomics (Dahlberg et al. 2009). From the ecological point of view, their relatively short life cycle, their ability to thrive in eutrophic (nutrient-rich) environments and a lack of significant predators contribute to their success in newly introduced environments (Lambert 2001 Shenkar and Loya 2008). The corollary is that ascidians are among the worst marine invasive species and that their rate of introduction has increased during the past decade (Lambert 2009). It is thus essential to develop tools that enable us to distinguish nonindigenous from indigenous species and ascertain the source populations of the introduced species. Unfortunately, ascidian systematics is notoriously difficult, because species are mostly classified based on inner anatomical characters such as gonad or gut loop shape and positions, and branchial sac structures (Monniot et al. 1991). Consequently, misidentifications of ascidian species are frequent (Mastrototaro and Dappiano 2008 Lambert 2009). Molecular sequences provide a way to complement species identification, especially in situations where traditional morphology-based discrimination of taxa is inadequate (Geller et al. 2010). Molecular markers, and in particular mitochondrial (mt) DNA sequences, thus provide a powerful alternative to the morphological approach. As a case in point, mt DNA has been successfully used to unequivocally demonstrate the existence of two cryptic species in the cosmopolitan ascidian Ciona intestinalis (Iannelli, Pesole, et al. 2007). Notwithstanding, ascidians are fast-evolving species (Yokobori et al. 1999, 2005 Tsagkogeorga, Turon, et al. 2010), a feature that complicates the use of their molecular characters to infer their evolutionary history (Delsuc et al. 2006). More specifically, ascidian mt genomes are hypervariable in almost all genomic features, which include for example, extremely high rates of sequence divergence and rampant gene order rearrangements, even at low taxonomic levels such as in congeneric and cryptic species (Iannelli, Griggio, et al. 2007 Gissi et al. 2010). This extremely fast evolution of ascidian mt genomes makes their sequence amplification a challenging task, which in turn explains the paucity of these sequenced genomes. We thus aimed to develop a simple and efficient method by which complete ascidian mt genomes can be easily acquired.

      Next-generation sequencing (NGS) technologies have revolutionized data acquisition in biology. Although sequencing protocols were originally developed for extracting a genome or transcriptome from a single organism, it is possible to mix several samples in a single flow cell (i.e., multiplex sequencing) as long as the sequences from the different samples can be subsequently separated. Standard multiplex methods allow pooling up to 96 different samples by introducing barcodes (or tags) during the DNA library preparation (Binladen et al. 2007). Following the sequencing step, reads are separated based on their barcode tags, such that assembly is performed for each sample separately. The advantage of this approach is the possibility to establish a trade-off between the total number of reads available from a single NGS run and the number of reads required to obtain a desired coverage for each individual sample. However, the disadvantage of such an approach is that it requires constructing separate genomic libraries for each sample, which can be costly. Several studies have suggested mixing several samples without barcoding them and separating the sequences only after the assembly step (Pollock et al. 2000 McComish et al. 2010 Timmermans et al. 2010 Dettai et al. 2012). We refer here only to nontheoretical studies. In Timmermans et al. (2010), the postassembly separation was based on bait sequences, which are short sequences (200𠄱,000 bp) obtained for each sample using Sanger sequencing. In McComish et al. (2010), the separation was performed by comparing the assembled contigs to a set of closely related reference mt genomes. Both Timmermans et al. (2010) and McComish et al. (2010) sequenced long polymerase chain reaction (PCR) amplified fragments covering the entire mitogenome. Unfortunately, the acquisition of long PCR fragments is extremely difficult in tunicates due to the pervasive gene order rearrangements. In addition, PCR artifacts can sometimes give rise to chimeric mt contigs (Timmermans et al. 2010).

      In this work, we chose to use the Illumina platform to sequence total genomic extracts of multiple species mixed together. Thus, both nuclear and mt DNA fragments of multiple species were sequenced together, and the mtDNA sequences were computationally retrieved through the assembly step. Our approach is similar to that used by Groenenberg et al. (2012), who obtained the complete mitogenome of a snail by Illumina sequencing and de novo assembly of the total DNA extracted from a single museum specimen. Following Timmermans et al. (2010), bait sequences were here used to identify the mt sequences of each sample rather than closely related sequences, as in McComish et al. (2010), since we sequenced, for example, the first representative of a family whose phylogenetic position is debated (e.g., Corellidae Tsagkogeorga et al. 2009). The advantage of our 𠇋rute force” approach is that it neither depends on specific primers nor on enrichment protocols and it is blind to mt gene order. Using this approach, we successfully assembled five new complete mitogenomes: Rhodosoma turcicum (Phlebobranchia: Corellidae), Botrylloides aff. leachii and Polycarpa mytiligera (Stolidobranchia: Styelidae), Halocynthia spinosa, and Pyura gangelion (Stolidobranchia: Pyuridae) ( fig. 1 A and CF, respectively). In addition, using the standard PCR and Sanger sequencing approaches, we obtained the mt genome of Ascidiella aspersa (Phlebobranchia: Ascidiidae) ( fig. 1 B). We describe and discuss our novel approach to mtDNA sequencing using NGS technology, together with the characteristics of these six new ascidian mt genomes in terms of genome organization and phylogenetic signal.

      Ascidian species sequenced in this work. (A) Rhodosoma turcicum (Corellidae), (B) Ascidiella aspersa (Ascidiidae), (C) Botrylloides aff. leachii (Styelidae), (D) Polycarpa mytiligera (Styelidae), (E) Halocynthia spinosa (Pyuridae), and (F) Pyura gangelion (Pyuridae).


      Purification and Concentration of DNA from Aqueous Solutions

      This unit presents basic procedures for manipulating solutions of single- or double-stranded DNA through purification and concentration steps. These techniques are useful when proteins or solute molecules need to be removed from aqueous solutions, or when DNA solutions need to be concentrated. The

      , using phenol extraction and ethanol (or isopropanol) precipitation, is appropriate for purification of DNA from small volumes (<0.4 ml) at concentrations lower than 1 mg/ml. Three support protocols outline methods to buffer the phenol used in extractions, concentrate DNA using butanol, and extract residual organic solvents with ether. An alternative to these methods is nucleic acid purification using glass beads and this is also presented. These protocols may also be used for purifying RNA. The final two alternate protocols are used for concentrating RNA and extracting and precipitating DNA from larger volumes and from dilute solutions, and for removing low-molecular-weight oligonucleotides and triphosphates.


      DNA Extraction from Plant Leaves Using a Microneedle Patch

      Isolation of high-quality DNA from infected plant specimens is an essential step for the molecular detection of plant pathogens. However, DNA isolation from plant cells surrounded by rigid polysaccharide cell walls involves complicated steps and requires benchtop laboratory equipment. As a result, plant DNA extraction is currently confined to well-equipped laboratories and sample preparation has become one of the major hurdles for on-site molecular detection of plant pathogens. To overcome this hurdle, a simple DNA extraction method from plant leaf tissues has been developed. A microneedle (MN) patch made of polyvinyl alcohol (PVA) can isolate plant or pathogenic DNA from different plant species within a minute. During DNA extraction, the polymeric MN patch penetrates into plant leaf tissues and breaks rigid plant cell walls to isolate intracellular DNA. The extracted DNA is polymerase chain reaction (PCR) amplifiable without additional purification. This minimally invasive method has successfully extracted Phytophthora infestans DNA from infected tomato leaves. Moreover, the MN patch could be used to isolate DNA from other plant pathogens directly in the field. Thus, it has great potential to become a rapid, on-site sample preparation technique for plant pathogen detection. © 2020 by John Wiley & Sons, Inc.

      Basic Protocol: Microneedle patch-based DNA extraction

      Support Protocol 1: Microneedle patch fabrication

      Support Protocol 2: Real-time PCR amplification of microneedle patch extracted DNA


      References

      Sinha R, Abu-Ali G, Vogtmann E, Fodor AA, Ren B, Amir A, Schwager E, Crabtree J, Ma S, Abnet CC, et al. Assessment of variation in microbial community amplicon sequencing by the Microbiome Quality Control (MBQC) project consortium. Nat Biotechnol. 201735:1077–86.

      Costea PI, Zeller G, Sunagawa S, Pelletier E, Alberti A, Levenez F, Tramontano M, Driessen M, Hercog R, Jung FE, et al. Towards standards for human fecal sample processing in metagenomic studies. Nat Biotechnol. 201735:1069–76.

      The Human Microbiome Project Consortium. Structure, function and diversity of the healthy human microbiome. Nature. 2012486:207–14.

      Qin J, Li R, Raes J, Arumugam M, Burgdorf KS, Manichanh C, Nielsen T, Pons N, Levenez F, Yamada T, et al. A human gut microbial gene catalogue established by metagenomic sequencing. Nature. 2010464:59–65.

      Marotz C, Amir A, Humphrey G, Gaffney J, Gogul G, Knight R. DNA extraction for streamlined metagenomics of diverse environmental samples. Biotechniques. 201762:290–3.

      Wesolowska-Andersen A, Bahl MI, Carvalho V, Kristiansen K, Sicheritz-Ponten T, Gupta R, Licht TR. Choice of bacterial DNA extraction method from fecal material influences community structure as evaluated by metagenomic analysis. Microbiome. 20142:19.

      Franzosa EA, Morgan XC, Segata N, Waldron L, Reyes J, Earl AM, Giannoukos G, Boylan MR, Ciulla D, Gevers D, et al. Relating the metatranscriptome and metagenome of the human gut. Proc Natl Acad Sci U S A. 2014111:E2329–38.

      Horz HP, Scheer S, Huenger F, Vianna ME, Conrads G. Selective isolation of bacterial DNA from human clinical specimens. J Microbiol Methods. 200872:98–102.

      Marotz CA, Sanders JG, Zuniga C, Zaramela LS, Knight R, Zengler K. Improving saliva shotgun metagenomics by chemical host DNA depletion. Microbiome. 20186:42.

      Eisenhofer R, Minich JJ, Marotz C, Cooper A, Knight R, Weyrich LS. Contamination in low microbial biomass microbiome studies: issues and recommendations. Trends Microbiol. 201927:105–17.

      Salter SJ, Cox MJ, Turek EM, Calus ST, Cookson WO, Moffatt MF, Turner P, Parkhill J, Loman NJ, Walker AW. Reagent and laboratory contamination can critically impact sequence-based microbiome analyses. BMC Biol. 201412:87.

      Glassing A, Dowd SE, Galandiuk S, Davis B, Chiodini RJ. Inherent bacterial DNA contamination of extraction and sequencing reagents may affect interpretation of microbiota in low bacterial biomass samples. Gut Pathog. 20168:24.

      Minich JJ, Zhu Q, Janssen S, Hendrickson R, Amir A, Vetter R, Hyde J, Doty MM, Stillwell K, Benardini J, et al. KatharoSeq enables high-throughput microbiome analysis from low-biomass samples. mSystems. 20183.

      Morales E, Chen J, Greathouse KL. Compositional analysis of the human microbiome in cancer research. Methods Mol Biol. 19282019:299–335.

      Dejea CM, Wick EC, Hechenbleikner EM, White JR, Mark Welch JL, Rossetti BJ, Peterson SN, Snesrud EC, Borisy GG, Lazarev M, et al. Microbiota organization is a distinct feature of proximal colorectal cancers. Proc Natl Acad Sci U S A. 2014111:18321–6.

      Bullman S, Pedamallu CS, Sicinska E, Clancy TE, Zhang X, Cai D, Neuberg D, Huang K, Guevara F, Nelson T, et al. Analysis of Fusobacterium persistence and antibiotic response in colorectal cancer. Science. 2017358:1443–8.

      Huseyin CE, Rubio RC, O'Sullivan O, Cotter PD, Scanlan PD. The fungal frontier: a comparative analysis of methods used in the study of the human gut mycobiome. Front Microbiol. 20178:1432.

      Rosenbaum J, Usyk M, Chen Z, Zolnik CP, Jones HE, Waldron L, Dowd JB, Thorpe LE, Burk RD. Evaluation of oral cavity DNA extraction methods on bacterial and fungal microbiota. Sci Rep. 20199:1531.

      Shkoporov AN, Ryan FJ, Draper LA, Forde A, Stockdale SR, Daly KM, McDonnell SA, Nolan JA, Sutton TDS, Dalmasso M, et al. Reproducible protocols for metagenomic analysis of human faecal phageomes. Microbiome. 20186:68.

      Kim D, Hofstaedter CE, Zhao C, Mattei L, Tanes C, Clarke E, Lauder A, Sherrill-Mix S, Chehoud C, Kelsen J, et al. Optimizing methods and dodging pitfalls in microbiome research. Microbiome. 20175:52.

      Hornung BVH, Zwittink RD, Kuijper EJ. Issues and current standards of controls in microbiome research. FEMS Microbiol Ecol. 201995. https://doi.org/10.1093/femsec/fiz045

      Sinha R, Ahsan H, Blaser M, Caporaso JG, Carmical JR, Chan AT, Fodor A, Gail MH, Harris CC, Helzlsouer K, et al: Next steps in studying the human microbiome and health in prospective studies, Bethesda, MD, May 16-17, 2017. Microbiome 2018, 6:210.

      Sinha R, Abnet CC, White O, Knight R, Huttenhower C. The microbiome quality control project: baseline study design and future directions. Genome Biol. 201516:276.


      Biology Lab Report on the extraction of Chlorophyl

      Abstract This experiment focused on extracting and separating pigments of Chloroplast. For the procedure green leaves were grinded in a mortar with some chemicals and the fluid was filtrated to use for further analysis. Stripes of this solution were put on a filter paper and later, after dried placed into a beacon of solvent. After this the chloroplast pigments were separated by the solvent into groups of more or less soluble pigments.

      Aim How many pigment types are present in a green leaf? It is hoped to be able to identify the four different pigments types of a leaf.

      As the filter paper with the solvent will separate the pigments in terms of solubility, a clear segmentation of each is expected to show off. As various chemicals were used in the whole process of this lab, certain variables might have influenced the results in terms of the purity of each chemical or purity of the used filtrate.

      Background The chloroplast, basically, is the organelle responsible for photosynthesis.

      Structurally it is very similar to the mitochondria. It contains a permeable outer membrane, a less permeable inner membrane, a intermediate space, and an inner section called the storms. However, the chloroplast is larger than the mitochondria. It needs to have the larger size because its membrane is not folded into Cristal. Also the inner membrane is not used for the electron transport chain.

      In fact, the first filtrate we used wasn’t even containing enough pigments which reacted to the solvent to create any useful results. After redoing the methods for the filtrate and paper strip the solvent started to do its Job. Within about 10 minutes the solvent had slowly sucked up into the paper till the grey pencil line. Then, after having dried the strip one could very scarcely identify two distinct colors, each only a couple of millimeters away of each there: a bluish green and a yellowish green line.

      Teacher Glen then gave us a scale upon which we should identify each pigment strip with its individual color, measure its distance in relation to the origin (the first line drawn with the filtrate) and then calculate the REF. First one had to identify the precise color of each, this was a bit problematic, due to the gamut of different tones present. As presented in figure 1 or Data Table 2, there is a set order of the distribution of each color band. The REF was calculated upon the following formula: REF = Distance Pigment migrated (mm)

      Distance solvent front migrated (mm) Figure 1 – Carotene (orange) – Carotene – Psychoanalyst Pigments in Chloroplast – Chlorophyll – Chlorophyll A (bluish green) – Chlorophyll B (yellowish green) Conclusion As expected the solvent has caused the pigments to migrate into their groups of solubility. The yellowish green Chlorophyll B was the first band we identified, only migrated mm and the bluish green Chlorophyll A migrated mm. This told us that there was only Chlorophyll present in the green leave, although other students who had performed this experiment as well achieved different results.

      The only issue which could be mentioned about the overall lab design is the rooms’ ventilation system. After having opened the bottle with the solvent, within only a couple of minutes the whole room had a very intense smell which made a few students feel dizzy or get a headache. There are no real suggestions for this problem, except talking to the schools maintenance crew and tell them to get the fans fixed. The reasons for the strange results could only be derived due to the fact, that maybe the chemicals used for this experiment weren’t pure enough, as well as the self made filtrate.


      Watch the video: 02 - DNA Isolation from bacterial culture (October 2022).