Video Capture and Editing: Protocols and Applications

This section is an expanded web version of the methods described in the following papers:

De Ley, P., Bert, W. 2002. Video Capture and Editing as a tool for the storage, distribution and illustration of morphological characters of nematodes. Journal of Nematology. Vol. 34: p.296-302.

De Ley, P., De Ley, I.T., Morris, K., Abebe, E., Mundo-Ocampo, M., Yoder, M., Heras, J., Waumann, D., Rocha-Olivares, A., Burr, A.J., Baldwin, J.G., Thomas, W.K. 2005. An integrated approach to fast and informative morphological vouchering of nematodes for applications in molecular barcoding. Philosophical Transactions of the Royal Society of London B. Vol. 360: p.1945-1958.

Yoder, M., Tandingan De Ley, I., King, I.W., Mundo-Ocampo, M., Mann, J., Blaxter, M., Poiras, L., De Ley, P. 2006. DESS: a versatile solution for preserving morphology and extractable DNA of nematodes. Nematology. Vol. 8: p.367-376.

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Material and Methods

Summary: Morphological identification of nematodes usually requires permanent slides for detailed observation. This significantly limits the range of research methods that can be applied to such specimens. Permanent slides are never truly permanent, as taxonomically important reference material may be lost by accident, or degrades slowly over time. In order to efficiently record the morphology of nematodes, in a format that allows easy archiving, editing and distribution, we have assembled and tested two micrographic Video Capture and Editing (VCE) configurations. These comparatively inexpensive, easily customized systems allow for the production of short video clips that mimic multifocal observation of nematode specimens through a light microscope. These clips can be used for many purposes, including teaching and training, management and online access of taxonomic collections, routine screening of fixed or unfixed specimens, recording of ephemeral staining patterns, or recording of freshly dissected internal organs prior to their decomposition. We provide details on the components and operation of both systems, evaluate their efficiency in the aforementioned applications, and provide illustrations of the obtained image quality. We expect this approach to become widely used in nematology research and teaching.

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Most nematodes are transparent micro-invertebrates, and the light microscope (LM) is therefore the first-line tool for nematode identification. This will undoubtedly remain true for a long time, despite the proliferation in recent decades of various ultrastructural and molecular techniques. Unfortunately, many situations occur in which a specimen's micro-morphological characters are irreversibly lost. For example, type specimens deposited in taxonomic reference collections deteriorate inevitably, and are sometimes accidentally lost or damaged, while teaching material often gets destroyed by inexperienced hands long before preservation fails. Specimens prepared for histochemical studies or Scanning Electron Microscopy (SEM) lose transparency during staining or coating protocols, and often deteriorate within hours, days or weeks. Certain kinds of observations on internal structures, such as the cellular architecture of the female gonad, can only be made on freshly dissected specimens, which usually plasmolyse and decompose quickly. Even more drastically, techniques such as Transmission Electron Microscopy (TEM) and molecular analysis require the destruction of the nematode being studied, as a prerequisite to the extraction of ultrastructural or macromolecular data.

Loss of optical morphology of a specimen invariably interferes with subsequent verification of previous identifications and observations. It also complicates procedures for applying multiple methodologies to the same individual nematode. For example, although it is possible to obtain DNA sequence data from single nematodes fixed years earlier in formalin (Dorris & Blaxter, pers. comm.), chances of success are highest when DNA extraction is performed soon after fixation (Thomas et al., 1997) in order to maximize succesful PCR amplification. And regardless of the time elapsed since fixation, at least part of the specimen must inevitably be destroyed. Some of the consequences are, (i) that morphology-based identification of an individual nematode must essentially be complete before its molecular analysis can begin, (ii) that curators of reference collections can rarely permit attempts at molecular analysis of older type specimens, and (iii) that it constrains practical feasibility of large-scale surveys of nematode diversity combining morphological and molecular data of each individual.

For many years, line drawings and still photographs have acted as the sole means of illustrating, distributing and preserving LM-visible morphology. Both types of images have important drawbacks. First, they depend on the skills of the person producing them (especially so for line drawings), and second, they simplify or omit three-dimensional topology of structures (especially so for still photos). Eisenback (1988) presented a simple method for superimposing several focal planes in one photomicrograph, but this method is only applicable to a maximum of five focal planes, and the act of superimposition reduces contrast per focal plane, as well as discarding all topological information along the focal axis.

We have therefore set out to find alternative approaches for recording the morphological characters of nematodes as seen with LM. Taking cues from four-dimensional microscopy, as utilized in comparative embryology of nematodes for multifocal image recording through time (Thomas et al., 1996; Schnabel et al., 1997), we have assembled two simplified, non-automated configurations for multifocal recording of nematode morphology. We assembled both general-purpose Video Capture and Editing (VCE) systems from inexpensive and widely available components, tested them and developed simple protocols for the recording, optimization and distribution of multifocal images of nematode specimens. The resulting video files can easily be exchanged on disk or through networks, and can be viewed with various widely distributed video player programs.

In this paper we describe the various components and protocols used for our VCE installations, as well as their relevant functions, problems and observed disadvantages. We conclude that VCE has a wide range of possible uses in nematological teaching and research, and that it is complementary to still photography rather than a direct methodological competitor or replacement.

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