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locktite seeds

Locktite seeds

ALSO, you are still responsible for controling the noxious weeds on your property.

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Provided is a form below is a form on the bottom of the page to print and send to the weed district. If preferable you may also send an email to the weed district : [email protected]


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Locktite seeds

The resin blocks with the specimens adequately embedded are then cut into serial thin sections, or wafers, using a lapidary low-speed saw with a diamond cutting edge (e.g., IsoMet Low Speed Saw; Buehler, Lake Bluff, Illinois, USA), which uniformly cuts through the specimen and resin to yield wafers 0.5–1.0 mm thick, depending on the type of blade and lapidary saw used. The wafers are then cleaned, dried, and affixed to a traditional microscope slide using any standard mountant typically used to affix fossil wafers to slides (e.g., Manchester, 1994 [Elmer’s epoxy; Elmer’s Products Inc., Westerville, Ohio, USA]; Pigg et al., 2004 [UV-cured adhesive, UV-154; T.H.E. Company, Lakewood, Colorado, USA]; Rothwell and Ash, 2015 [Devcon 5 Minute Epoxy; ITW Devcon, Danvers, Massachusetts, USA]). The specimens are then photographed to document general morphology and three-dimensional features that could be lost when grinding specimens for transmitted microscopy. The specimens are then taken through a series of coarse- to fine-grit sandpaper or polishing papers (e.g., 3M Wetordry 6-pc assorted 1–30-μm aluminum oxide/silicon carbide polishing papers; 3M, St. Paul, Minnesota, USA) and ground down to a minimal thickness (<100 μm). Upon completing grinding and polishing, specimens can be made into permanent slides by adhering a coverslip with a long-term mounting medium (e.g., Eukitt; Sigma-Aldrich, St. Louis, Missouri, USA; see Ruzin [1999] for a list of common mounting media or Brown [1997] for a review of various media) and then photographed using transmitted light microscopy ( Fig. 2A–H ). The author has prepared a variety of plant specimens in this manner for approximately 10 yr and no deformation of tissues as a result of dehydration has been observed; these slides should last as long as traditionally prepared microscope slides.

In some seeds, the embryo or nutrient tissue may loosen during the cutting process. To avoid specimen loss, apply a thin layer of nail polish onto the cutting surface and allow it to dry prior to each cut. This will hold the various tissues in place during the cutting process. The nail polish can then be removed using acetone or another nail polish remover after the thin section has been cut. Number each thin section, remove oil with soapy water or ethanol, and select optimal cuts to be mounted on microscope slides. A water-based saw may also be used (e.g., Hi-Tech Diamond 6″ Trim Saw), but specimens should not be left in water long because fibers and sclereids may become disfigured and warp the wafer. Both sides of a section should be analyzed and the optimal side noted prior to mounting. Also, it is wise to take overview photographs of the specimens at this stage to document the overall morphology of the specimen (e.g., Figs. 1A–B , ​ ,2D) 2D ) and to determine how much grinding is needed to obtain roughly a single layer of cells for analysis.


Many methods exist for producing thin sections of various plant tissues for anatomical studies, but few methods exist for highly sclerified and heterogeneous tissues of fruits and seeds. The technique outlined here effectively prepares fruit and seed tissues from a variety of gymnosperm and angiosperm families for which traditional embedding/sectioning methods have failed. It has the advantage of dramatically decreasing the processing time of materials from weeks to months, to a maximum of three days. It eliminates the need for polar and nonpolar solvents, which leaves the chemical composition of materials processed intact. It also avoids tearing hard or brittle tissues by removing the need for a sharp blade to section material, which can damage the sectioning apparatus or the desired tissues.

Fruit and seed morphology and anatomy using the thin sectioning technique. (A–C) Longitudinal section of Alpinia malaccensis (Zingiberaceae). (A) Overview of longitudinal section of the seed before grinding. (B) Detail of the operculum and micropylar region of the seed before grinding. (C) Detail of the seed coat after grinding with a yellow, palisade exotesta, red to yellow mesotesta, and dark red sclerenchymatous endotesta. (D–F) Transverse section of a maple fruit (Acer platanoides; Sapindaceae). (D) Overview of a maple fruit in transverse section before grinding. (E) Detail of the fruit wall after grinding. (F) Detail of the fruit wall (at bottom), seed coat (red, middle), and embryo (top) after grinding. (G) Detail of the seed coat of Distylium racemosum (Hamamelidaceae) after grinding. (H) Detail of the seed coat of Hamamelis virginiana (Hamamelidaceae) after grinding. Scale bars: A, D = 1 mm; B = 500 μm; C = 50 μm; E–H = 100 μm.


When compared to existing techniques for obtaining anatomical features of fruits and seeds, the protocol described here has the potential to create high-quality thin sections of materials that are not able to be sectioned using traditional histological techniques, which can be produced quickly and without the need for harmful chemicals.