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Techniques for Viewing Pollen Tubes in Angiosperm Flowers Cameron Thompson [email protected]

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Techniques for Viewing Pollen Tubes in Angiosperm Flowers Cameron Thompson Cdt40@Zips.Uakron.Edu The University of Akron IdeaExchange@UAkron The Dr. Gary B. and Pamela S. Williams Honors Honors Research Projects College Spring 2016 Techniques for Viewing Pollen Tubes in Angiosperm Flowers Cameron Thompson [email protected] Please take a moment to share how this work helps you through this survey. Your feedback will be important as we plan further development of our repository. Follow this and additional works at: http://ideaexchange.uakron.edu/honors_research_projects Recommended Citation Thompson, Cameron, "Techniques for Viewing Pollen Tubes in Angiosperm Flowers" (2016). Honors Research Projects. 390. http://ideaexchange.uakron.edu/honors_research_projects/390 This Honors Research Project is brought to you for free and open access by The Dr. Gary B. and Pamela S. Williams Honors College at IdeaExchange@UAkron, the institutional repository of The nivU ersity of Akron in Akron, Ohio, USA. It has been accepted for inclusion in Honors Research Projects by an authorized administrator of IdeaExchange@UAkron. For more information, please contact [email protected], [email protected]. Techniques for Viewing Pollen Tubes in Angiosperm Flowers Thompson, C., Mitchell, R. Abstract: In angiosperm plants, pollen is produced in the anthers and dispersed by pollinators to the stigmas of either the same plant or other flowers. After pollen reaches the stigma, pollen tubes grow down the style transporting the male gametes, cytoplasm, and a vegetative nucleus to the ovary, which fertilizes the flower. Currently there are many different techniques to stain and study pollen tubes in order to investigate reproductive techniques of many angiosperms. Though many techniques are available, few result in images that facilitate counting of pollen tubes at different stages of pollination. In this study, a number of promising techniques for fixing, softening, and staining stigmas were compared to determine the most efficient way of viewing the numerous pollen tubes in terms of time, cost, and images produced, and therefore learn more about the reproduction process in angiosperms. Introduction different points along the style could lead to a better understanding of In angiosperms, including Monkey pollen tube development including flowers of the genus Mimulus, the which factors are most critical to male gametes are produced in pollen effective pollination. grains on the anther of the flower. Pollinators, including bees and birds, Staining is a technique used in while harvesting from the flower, get histology to facilitate observing pollen on their bodies, which they different structures, tissues, or disperse to other flowers. Once the organisms through artificial pollen is on the stigma of the flower, it coloration. The staining process uses the nutrients stored inside the consists of three main steps, including pollen grain, and begins to extend in fixing the object, softening it, and then the form of a pollen tube towards the staining it (Kearns and Inouye, 1993). female gamete in the carpel In this experiment, fixing agents are (Krichevsky, et al, 2006). After used to cease growth of the pollen traveling approximately 1/3 of the tubes and allow for the storage of the way towards the female gamete, the samples for long periods of time pollen grain must then use nutrients without damaging the sample. from the stigma to travel the Common fixing solutions were used in remaining length. While the pollen this study and included ethyl alcohol, tubes travel towards the ovary as a Farmer’s solution, and formalin-acetic vehicle for the cytoplasm, nucleus, and acid-alcohol (FAA) solution. Sodium male gametes, deposits of callose are sulphite and different concentrations periodically placed to seal the tube of sodium hydroxide were tested as into segments (Qin, et al, 2012). Being softening agents, which soften the able to count the pollen tubes at tissues and make them more receptive to dyes. In plant histology, the most was placed in either: 70% ethyl commonly used method for the alcohol, Farmer’s solution prepared staining of pollen tubes is the aniline from 3 parts ethyl alcohol to 1 part blue method, which stains the callose acetic acid, or formalin-acetic acid- plugs of the tubes, which then become alcohol (FAA) solution. The FAA viewable under a fluorescent light solution was prepared using 10 microscope. Though the method milliliters of formalin, 5 milliliters of works well, not all plants deposit an acetic acid, 50 ml of ethanol, and 35 ml abundance of callose, and the aniline of deionized water. The samples fixed blue technique does not facilitate easy in farmer’s solution and the FAA counting of pollen tubes in Mimulus solution were then transferred to 70% because the deposits are long EtOH after 24 hours. After fixing for at intervals. Other methods for staining least 24 hours, the fixing agent was were used in this experiment to see if removed and replaced by distilled the callose plugs could be made water for 10 minutes, which was in brighter or clearer, or other aspects of turn removed and replaced by a the pollen tube could be stained to softening solution. To soften the facilitate imaging and counting. These tissues, a fraction of pistils from each other methods included Alexander’s fixing solution were then softened combination of four stains, staining with one of the following solutions: a with acidified aniline blue and 0.4M sodium sulphite solution, a 1 acetocarmine, a combination of molar sodium hydroxide, an 8 M acetocarmine and basic fuchsin stains, NaOH, or a 10 M NaOH for a period of the use of a fluorescent brightener in 4-24 hours. After softening, the tissues conjunction with aniline blue, and were washed again with distilled Congo red staining. water in their bullet tubes, their ovaries were removed, and samples Methods from each group were stained with a one of the following methods of Samples of Mimulus ringens flowers staining: aniline blue, aniline blue with were collected throughout the fluorescent brightener, acetocarmine flowering season, from the Panzner combined with acidified aniline blue, Wetlands in Akron, Ohio, where they acetocarmine with basic fuchsin, grow naturally. Flowers were Alexander’s combination of four collected during the months of July stains, or Congo red. and August in 2015 between the hours of 11 am and 5 pm. Samples The Aniline blue stain was adapted consisted of pistils both with and from the protocol outlined by Mori, T. without petals. Pistils were harvested in Generative Cell Specific 1 is Essential by removing petals, if applicable, and for Angiosperm Fertilization. After sepals, placed in individual bullet fixing and softening using previously tubes and stored in a fixing agent stated techniques, stigmas were within 24-48 hours. Samples not decolorized using a solution of 0.1% immediately fixed were stored in a aniline blue in potassium phosphate refrigerator at approximately 4 for 2 hours in dark conditions. After degrees Celsius. For fixing, each pistil decolorization, stigmas were mounted on slides with a drop of decolorized Pollination Biologists. 1% solutions of aniline blue prior to mounting the malachite green, acid fuchsin, and coverslip. aniline blue in distilled water, and a 1% solution of Orange G in 50% ethyl Acidified aniline blue was prepared alcohol were prepared for use. Then using a 0.1% aniline blue solution in 78 ml of lactic acid, 4 ml of the K3PO4 with the addition of malachite green solution, 6 ml of the concentrated hydrochloric acid until acid fuchsin solution, 4 ml of the the solution was acidic to litmus aniline blue solution, and 2 ml of the paper. A solution of acetocarmine was orange G solution were combined in a prepared by saturating a carmine light resistant jar. Stigmas were fixed, powder with 40% acetic acid. After transferred to the combination stain, fixing and softening, stigmas were then softened and mounted in a 1:1 mounted on a slide with a drop of the medium of lactic acid and glycerol. combined stain. A 0.1% Congo red solution in distilled Another subsection of the samples water was prepared as a stain for the was treated with a drop of the final technique. Tissues were fixed aforementioned acetocarmine and softened using the previous solution followed by a drop of 3% methods and a drop of the stain was basic fuchsin in distilled water. Excess placed on each sample. The samples stain was removed after 10 minutes were then mounted in either another and a drop of ethyl alcohol was added drop of stain or in glycerin. to each sample and then absorbed away with filter paper. Samples were All slides were then viewed and then mounted using glycerin. imaged with a Zeiss Axioplan 2 microscope with both halogen light A solution including the fluorescent and fluorescence and evaluated for brightener calcofluor was used to visibility and accessibility of pollen stain another portion of stigmas. The tubes. solution was prepared by dissolving 0.5 grams of aniline blue, 0.5 grams of Results brightener, and 1.75 grams of K3PO4 individually in small amounts of Though all three fixing methods distilled water, and then combined, worked efficiently, using Farmer’s with the total volume being brought solution or FAA provided a up to 500 ml. After fixing and consistently clearer image of the softening the tissues, a drop of the subject based on images with the combined stain was placed on the same softening and staining but stigmas, and after 10 minutes, the different fixation procedures. 70% stigmas were placed in wet mounts for EtOH appeared to not halt organic viewing. functions fast enough, resulting in the pistils decomposing, while Farmer’s Alexander’s combination of four stains solution appeared to preserve styles was prepared following a modified most effectively, with the pistils’ procedure outlined in Techniques for appearance remaining unchanged Both combination stains including throughout fixation.
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