**Ultrastructure and Topochemistry of Plant Cell Wall by Transmission Electron Microscopy**

Xia Zhou, Dayong Ding, Jing Ma, Zhe Ji, Xun Zhang and Feng Xu

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/60752

#### **Abstract**

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Plant cell walls are typically described as complex macromolecular composites consisting of an ordered array of cellulose microfibrils embedded in a matrix of noncellulosic polysaccharides and lignin. Generally, the plant cell wall can be divided into three major layers: middle lamella, primary cell wall, and secondary cell wall. Investigation of plant cell walls is complicated by the heterogeneous and complex hierarchical structure, as well as variable chemical composition between different sublayers. Thus, a complete understanding of the ultrastructure of plant cell walls is necessary. Transmission electron microscopy (TEM) has proven to be a powerful tool in elucidating fine details of plant cell walls at nanoscale. The present chapter describes the layering structure and topochemistry of plant cell wall revealed by TEM.

**Keywords:** Plant cell wall, Transmission electron microscopy, Ultrastructure, Topo‐ chemistry

### **1. Introduction**

Determining the ultrastructural organization of plant cell walls represents one of the most challenging problems in plant biology. Although considerable progress has been made in understanding the basic organization and functions of plant cell wall components, due to the highly complex and dynamic nature of the plant cell wall, the variation in cell wall architecture of wood and gramineous species remains poorly understood. These structural features are

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associated with cell growth and morphogenesis, which are also crucial in determining the mechanical properties of plant cell walls [1, 2]. Given that one of the critical processing steps in biomass conversion involves systematic deconstruction of cell walls, this structural infor‐ mation is also pivotal for developing novel approaches to convert biomass into liquid biofuels. Therefore, a comprehensive investigation of the architecture of the plant cell wall will not only help us to understand the assembly and biosynthesis of the plant cell wall, but will also contribute to improving the efficiency of biomass deconstruction [3].

The plant cell wall is a layered construction composed mainly of stiff crystalline cellulose microfibrils (Mfs) embedded in an amorphous matrix of non-crystalline cellulose, hemicellu‐ loses and pectin, as well as various aromatic compounds and proteins [4]. Besides the varieties of chemical constituents, the ultrastructural organization of plant cell wall varies between species and cell types. Generally, the plant cell wall consists of three major layers: (i) the middle lamella (Ml), (ii) the primary wall (Pw), and (iii) the secondary wall (Sw). Due to the highest thickness, Sw accounts for the largest proportion of the plant cell wall. Sw in higher plants consists mainly of cellulose, lignin, and xylan and is the major component of biomass in many species. In hardwood fibers and softwood tracheids, the Sw is normally further differentiated into an outer layer (S1), a middle layer (S2), and an inner layer (S3), with the S2 having the largest thickness [5, 6]. By comparison, in gramineous species the lamellation of the Sw in fiber is generally described as alternating broad and narrow layers [7].

In the twentieth century, microscopic approaches began to offer high-resolution images to enhance our understanding of cell wall organization. Over the past few decades, atomic force microscopy (AFM) has been successfully applied to high-resolution architecture, assembly, and structure dynamic studies of a wide range of biological systems, which has enabled researchers to visualize the ultrastructure of the plant cell wall [8, 9]. More recently, confocal Raman microspectroscopy (CRM) has now also been successfully applied to acquire informa‐ tion on the preferential orientation of plant polymer functional groups and components distribution in situ [10, 11]. However, although these approaches have been comprehensively used to obtain new information on cell wall architecture, until now the highly complex and dynamic nature of the plant cell wall at nanoscale has limited our ability to generate detailed structural models.

By comparison, due to the higher spatial resolution (<1nm) and specificity when combined with chemical staining and immunolabeling approaches, transmission electron microscopy (TEM) can provide ultrastructural and topochemical information simultaneously and has been used to investigate plant cell wall [12-14]. In this chapter, we mainly discuss the application of TEM in detecting cell wall layering structure and cell wall topochemistry (lignin distribution and hemicelluloses distribution).

### **2. Cell wall layering structure**

To get any information using transmitted electrons in the TEM, specimens have to be thin. "Thin" is a relative term, in this context it means electron transparent. For a specimen to be transparent to electrons, it must be thin enough to transmit sufficient electrons such that enough intensity falls on the screen, charge coupled device (CCD), or photographic plate to give an interpretable image in a reasonable time. Generally this requirement is a function of the electron energy and the average atomic number (Z) of the specimen. It is almost an axiom in TEM that thinner is better and specimens <100 nm should be used wherever possible. However, a too thin section would produce low-contrast TEM image, which hides the subtle structure. For plant cell wall, specimens are generally cut to a thickness of ~80 nm when they are silvery gold in color under ultramicrotome. In extreme cases such as doing high-resolution TEM (HRTEM) or electron spectrometry, specimen thicknesses <50 nm (even <10 nm) are essential.

### **2.1. Cell wall layering structure in hardwoods and softwoods**

associated with cell growth and morphogenesis, which are also crucial in determining the mechanical properties of plant cell walls [1, 2]. Given that one of the critical processing steps in biomass conversion involves systematic deconstruction of cell walls, this structural infor‐ mation is also pivotal for developing novel approaches to convert biomass into liquid biofuels. Therefore, a comprehensive investigation of the architecture of the plant cell wall will not only help us to understand the assembly and biosynthesis of the plant cell wall, but will also

The plant cell wall is a layered construction composed mainly of stiff crystalline cellulose microfibrils (Mfs) embedded in an amorphous matrix of non-crystalline cellulose, hemicellu‐ loses and pectin, as well as various aromatic compounds and proteins [4]. Besides the varieties of chemical constituents, the ultrastructural organization of plant cell wall varies between species and cell types. Generally, the plant cell wall consists of three major layers: (i) the middle lamella (Ml), (ii) the primary wall (Pw), and (iii) the secondary wall (Sw). Due to the highest thickness, Sw accounts for the largest proportion of the plant cell wall. Sw in higher plants consists mainly of cellulose, lignin, and xylan and is the major component of biomass in many species. In hardwood fibers and softwood tracheids, the Sw is normally further differentiated into an outer layer (S1), a middle layer (S2), and an inner layer (S3), with the S2 having the largest thickness [5, 6]. By comparison, in gramineous species the lamellation of the Sw in fiber

In the twentieth century, microscopic approaches began to offer high-resolution images to enhance our understanding of cell wall organization. Over the past few decades, atomic force microscopy (AFM) has been successfully applied to high-resolution architecture, assembly, and structure dynamic studies of a wide range of biological systems, which has enabled researchers to visualize the ultrastructure of the plant cell wall [8, 9]. More recently, confocal Raman microspectroscopy (CRM) has now also been successfully applied to acquire informa‐ tion on the preferential orientation of plant polymer functional groups and components distribution in situ [10, 11]. However, although these approaches have been comprehensively used to obtain new information on cell wall architecture, until now the highly complex and dynamic nature of the plant cell wall at nanoscale has limited our ability to generate detailed

By comparison, due to the higher spatial resolution (<1nm) and specificity when combined with chemical staining and immunolabeling approaches, transmission electron microscopy (TEM) can provide ultrastructural and topochemical information simultaneously and has been used to investigate plant cell wall [12-14]. In this chapter, we mainly discuss the application of TEM in detecting cell wall layering structure and cell wall topochemistry (lignin distribution

To get any information using transmitted electrons in the TEM, specimens have to be thin. "Thin" is a relative term, in this context it means electron transparent. For a specimen to be

contribute to improving the efficiency of biomass deconstruction [3].

286 The Transmission Electron Microscope – Theory and Applications

is generally described as alternating broad and narrow layers [7].

structural models.

and hemicelluloses distribution).

**2. Cell wall layering structure**

TEM examination showed that *Cornus alba* L*.* fiber cell wall was composed of three major layers: the middle lamella (ML) and the primary wall (P), and the secondary wall layer (S1, S2, and S3), a typical layering structure of fiber cell walls in other wood species [13, 15, 16]. The boundary between primary wall and middle lamella was not clearly distinguishable due to its high density and extreme thinness. Therefore, both the middle lamella (ML) and the contigu‐ ous primary wall (P) were referred here as compound middle lamella (CML) (Fig. 1a). The CML was electron dense, but the density was not uniform and this cell wall region had a mottled appearance containing dense and less dense or lucent regions. Inhomogeneity in lignin distribution in CML has also been reported in a few other TEM studies of hardwood species [17, 18]. The secondary wall was divided into an outer layer (F-S1), a middle layer (F-S2), and an inner layer (F-S3). The F-S1 layer in *Cornus alba* L. fiber was well-defined and can be readily distinguished from the adjoining F-S2 layer because of its higher electron density compared to F-S2 layer. The F-S1 layer was variable in width within and among cells (0.22–0.32 μm) (Table 1). The widest F-S2 layer accounted for the largest proportion of the fiber wall. Meas‐ urements of the width of fiber radial wall showed that the average thickness of the F-S2 layer was 2.67 μm. Fiber cell walls also contained an F-S3 layer that was very thin and not well developed.


**Table 1.** The average thickness of cell wall layers in *Cornus alba L.*

In addition to fiber cell wall, the ultrastructural variation in vessel, axial parenchyma, and ray parenchyma was also investigated. As shown in Fig. 1b, the vessel wall was divided into three layers (V-S1, V-S2, and V-S3) of variable electron density. The width of V-S1 ranged from 0.18 μm to 0.32 μm, approximately equal to that of F-S1, while the V-S2 was much thinner, with width from 0.55 μm to 0.67 μm. For the axial parenchyma (AP), the secondary wall was clearly resolved into an outer layer (AP-S1), a middle layer (AP-S2), and an inner layer (AP-S3) (Fig. 1c). Unlike the widest F-S2 layer accounting for the largest proportion of the secondary wall, in axial parenchyma the AP-S3 was the major proportion of secondary wall with the thickness ranging from 0.60 μm to 1.67 μm. In ray parenchyma (RP), the secondary wall consisted of two well-defined layers (outer layer, RP-S1, and inner layer, RP-S2), which did not fit conven‐ tional S1, S2, and S3 classification (Fig. 1d). Measurements taken on TEM micrograph evi‐ denced that the average width of the RP-S1 was 0.61 μm, while the average thickness of the RP-S2 was 0.80 μm.

**Figure 1.** TEM micrographs of cross sections of *C. alba* L., taken at 80 kV. (a) CCML, cell corner middle lamella between adjoining fibers; CML, compound middle lamella between adjoining fibers; F-S1, outer secondary wall of fiber; F-S2, middle secondary wall of fiber; F-S3, inner secondary wall of fiber. (b) V-S1, outer secondary wall of vessel; V-S2, mid‐ dle secondary wall of vessel; V-S3, inner secondary wall of vessel. (c) AP-S1, outer secondary wall of axial parenchy‐ ma; AP-S2, middle secondary wall of axial parenchyma; AP-S3, inner secondary wall of axial parenchyma. (d) RP-S1, outer secondary wall of ray parenchyma; RP-S2, inner secondary wall of ray parenchyma.

The ultrastructure of pit membrane (PM) among various wood elements (inter-fiber, fibervessel, fiber-axial parenchyma, and fiber-ray parenchyma) was also investigated. The thick‐ ness of PM varied considerably, with PM between fiber and ray parenchyma having a mean thickness of 500 nm, while PM between fiber and vessel had an average thickness of 220 nm. Thin PM with an average thickness of 230 nm was also found between parenchyma cells. PM varied also in their electron density, with inter-fiber PM (Fig. 2a) appearing distinctly denser than fiber-vessel (Fig. 2b) and fiber-parenchyma (axial and ray parenchyma) PM (Fig. 2c and 2d), which may reflect textural and/or compositional differences. The electron density variations originated from the deposition of lignin that is directly and linearly proportional to lignin concentration [13]. Thus, we can assume that the inter-fiber PMs have the highest lignin concentration, followed by fiber and parenchyma (axial and ray parenchyma) and fewest in the PM between fiber and vessel.

In addition to fiber cell wall, the ultrastructural variation in vessel, axial parenchyma, and ray parenchyma was also investigated. As shown in Fig. 1b, the vessel wall was divided into three layers (V-S1, V-S2, and V-S3) of variable electron density. The width of V-S1 ranged from 0.18 μm to 0.32 μm, approximately equal to that of F-S1, while the V-S2 was much thinner, with width from 0.55 μm to 0.67 μm. For the axial parenchyma (AP), the secondary wall was clearly resolved into an outer layer (AP-S1), a middle layer (AP-S2), and an inner layer (AP-S3) (Fig. 1c). Unlike the widest F-S2 layer accounting for the largest proportion of the secondary wall, in axial parenchyma the AP-S3 was the major proportion of secondary wall with the thickness ranging from 0.60 μm to 1.67 μm. In ray parenchyma (RP), the secondary wall consisted of two well-defined layers (outer layer, RP-S1, and inner layer, RP-S2), which did not fit conven‐ tional S1, S2, and S3 classification (Fig. 1d). Measurements taken on TEM micrograph evi‐ denced that the average width of the RP-S1 was 0.61 μm, while the average thickness of the

**Figure 1.** TEM micrographs of cross sections of *C. alba* L., taken at 80 kV. (a) CCML, cell corner middle lamella between adjoining fibers; CML, compound middle lamella between adjoining fibers; F-S1, outer secondary wall of fiber; F-S2, middle secondary wall of fiber; F-S3, inner secondary wall of fiber. (b) V-S1, outer secondary wall of vessel; V-S2, mid‐ dle secondary wall of vessel; V-S3, inner secondary wall of vessel. (c) AP-S1, outer secondary wall of axial parenchy‐ ma; AP-S2, middle secondary wall of axial parenchyma; AP-S3, inner secondary wall of axial parenchyma. (d) RP-S1,

outer secondary wall of ray parenchyma; RP-S2, inner secondary wall of ray parenchyma.

RP-S2 was 0.80 μm.

a b

288 The Transmission Electron Microscope – Theory and Applications

c d

**Figure 2.** TEM micrographs of pit membrane among various cells in *C. alba* L., taken at 80 kV. (a) Pit membrane be‐ tween fibers, (arrowheads: knife mark). (b) Pit membrane between fiber and vessel. (c) Pit membrane between fiber and ray parenchyma. (d) Pit membrane between fiber and axial parenchyma. F, fiber; V, vessel; RP, ray parenchyma; AP, axial parenchyma; PM, pit membrane.

Compared to hardwood, the cell type of softwood is uniform, mainly containing tracheids. In normal wood of *Pinus radiata* D. Don, the cell wall layering structure is similar to that of fiber in hardwood, while the structure of tracheid cell walls in compression wood is quite different [19]. TEM observations revealed a highly lignified outer S2 layer (S2L) and the absence of an S3 layer in compression wood of *Pinus radiata* D. Don (Fig. 3) [20].

**Figure 3.** TEM micrographs of mild compression wood of *Pinus radiata* D. Don*.*, taken at 80 kV. ML, middle lamella between adjoining tracheids; S1, outer secondary wall; S2, middle layer of secondary wall; S2L, outer S2 layer.

#### **2.2. Cell wall layering structure in gramineous species**

The investigation of the *Miscanthus sinensis* by TEM readily differentiated the sclerenchymat‐ ic fiber (Sf) into the middle lamella, the primary wall, and the secondary wall, a typical layering structure of fiber cell walls in wood and grass species (Fig. 4a) [13, 21, 22]. Interestingly, there was a greater degree of heterogeneity in the layering structure of Sf secondary wall (Fig. 4a and 4b), as represented by the fact that we were able to identify six main types (Type I-VI) depending on the number of alternating narrow and broad layers present. Narrow layers appeared as dark thin lines and had a more or less constant thickness. In order to establish the degree of variation in secondary wall patterning among fiber, a classification system of frequently observed patterns was devised based on TEM observations in Sf adjacent to xylem and phloem (Fig. 5). For the outer Sf connected to the surrounding parenchymatic tissue, the distribution of types I to III (4–6 layers) were predominant, whereas Type IV-VI (7-9 layers) were discernible in individual Sf close to the xylem vessel and phloem cells. Much variation in cell wall layering structure has also been reported for other bamboo species, such as *Phyllostachys viridiglaucescens* and *Dendrocala‐ mus asper*, in which the fibers are categorized into four and six major types according to their respective layering structure [21, 23]. Unlike the layering pattern of Sf adjacent to the xylem vessel in *Miscanthus sinensis*, *Dendrocalamus asper* fibers, which have the highest number of wall layers, were located at the periphery of the fiber bundles. Moreover, the poly-lamellated secondary wall is not an exclusive feature of herbaceous species and appears to have evolved in a variety of taxa. The sclerotic bark fibers of beech (*Fagus sylvatica*) have thick cell walls with numerous individual layers irregularly arranged [24]. Tension wood of *Laetia procera* (Poepp.) Eichl. (Flacourtiaceae) also shows a peculiar structure of secondary wall, which alters from thick to thin layers [25]. In the biomass conversion, the complex poly-lamellated structure acts like a barrier limiting the radial penetration of the chemicals and enzyme, which can be referred as the natural biomass recalcitrance.

in hardwood, while the structure of tracheid cell walls in compression wood is quite different [19]. TEM observations revealed a highly lignified outer S2 layer (S2L) and the absence of an

**Figure 3.** TEM micrographs of mild compression wood of *Pinus radiata* D. Don*.*, taken at 80 kV. ML, middle lamella between adjoining tracheids; S1, outer secondary wall; S2, middle layer of secondary wall; S2L, outer S2 layer.

The investigation of the *Miscanthus sinensis* by TEM readily differentiated the sclerenchymat‐ ic fiber (Sf) into the middle lamella, the primary wall, and the secondary wall, a typical layering structure of fiber cell walls in wood and grass species (Fig. 4a) [13, 21, 22]. Interestingly, there was a greater degree of heterogeneity in the layering structure of Sf secondary wall (Fig. 4a and 4b), as represented by the fact that we were able to identify six main types (Type I-VI) depending on the number of alternating narrow and broad layers present. Narrow layers appeared as dark thin lines and had a more or less constant thickness. In order to establish the degree of variation in secondary wall patterning among

**2.2. Cell wall layering structure in gramineous species**

S3 layer in compression wood of *Pinus radiata* D. Don (Fig. 3) [20].

290 The Transmission Electron Microscope – Theory and Applications

**Figure 4.** TEM images showing layering structure of Sf in *M. sinensis* cv. internode tissue, stained with 1% w/v KMnO4. (a) Sf adjacent to xylem; (b) Sf adjacent to phloem. Sf, sclerenchymatic fiber; Ccml, cell corner middle lamella; Cml, compound middle lamella; Sw, secondary wall.

In addition to the layering features of Sf secondary wall, the ultrastructural variation in conductive tissue (xylem vessels) was also visualized using TEM images (Fig. 6a and 6b). The secondary wall of Mxv and Pxv could not be clearly divided into sub-layers. This is probably due to either the uniform electron density or the cellulose microfibrils (Mfs) orientation.

**Figure 5.** Schematic illustration of Sf layering pattern. "Thin" layers are depicted as black lines, whereas the "broad" layers are colored green. A number has been assigned to the different types of layering for convenience. L: Lumen.

**Figure 6.** TEM images showing layering structure of *Miscanthus sinensis* internode tissue. (a) Metaxylem vessel (Mxv); (b) protoxylem vessel (Pxv).

### **3. Lignification and lignin distribution**

Next to cellulose, lignin is the most abundant and important polymeric organic substance in plant cell wall. It is a complex phenolic polymer formed by radical coupling reactions of three main monolignols: *p*-coumaryl, coniferyl, and sinapyl alcohol [26, 27]. As a major component of the cell wall of higher plants, lignin plays a vital role in plant growth by enhancing the strength of plant tissues and sealing the wall from water leaks and patho‐ gens invasion. During plant cell wall formation, lignification is generally regarded as the final stage of the differentiating process, where lignin is deposited within the polysacchar‐ ide cell wall framework by infilling interlamellar voids [28]. Studies of lignin distribution in plant cell walls are important because of the effect of uneven lignin distribution on wood properties, particularly pulping properties and resistance to decay. Until now, considera‐ ble effort has been applied to the investigation of lignin distribution in the cell wall. Since the early 1980s, advanced electron microscopic techniques were developed to obtain highresolution information on the lignin distribution in plant cell walls. TEM coupled with potassium permanganate (KMnO4) staining has proven to be effective in obtaining highresolution information on the lignin deposition [13, 19, 29]. On the other hand, the bromination technique by combination of energy-dispersive X-ray analysis (EDXA) with TEM or scanning electron microscopy (SEM) has shown the potential of providing quantitative information on the distribution of lignin. A similar method was developed with the mercurization of specimen and subsequent SEM/TEM-EDXA analyses, which is also based on chemical reactions between lignin and inorganic compounds [16, 30, 31]. Moreover, using newly developed immunological labeling markers to probe lignin with respect to its monomeric composition and with respect to the nature of its structural internal linkages, immunogold TEM has proven to be powerful in distinguishing different lignin substructures on the ultrastructural level [32-35].

#### **3.1. Lignification**

a b

292 The Transmission Electron Microscope – Theory and Applications

(b) protoxylem vessel (Pxv).

**Figure 6.** TEM images showing layering structure of *Miscanthus sinensis* internode tissue. (a) Metaxylem vessel (Mxv);

**Figure 5.** Schematic illustration of Sf layering pattern. "Thin" layers are depicted as black lines, whereas the "broad" layers are colored green. A number has been assigned to the different types of layering for convenience. L: Lumen.

> The lignification of plant cell walls is generally known to last for a long period, from the S1 stage to the F stage. After the enlargement of cell size, the secondary wall is thickened with the formation of the S1, S2, and S3 layers. The outermost region of the cell wall, including the intercellular layer, the cell comers, and the primary wall, is lignified during the S1 stage when the surface enlargement of the cell is completed, and just before the S1 starts thickening. This lignification, which will be called "intercellular layer (I)-lignification," may play an important role in stabilizing the cell size and adhering adjacent cells with one another. This I-lignification continues during the differentiation of the S1 and S2 layers, and even until the formation of the S3. On the other hand, the lignification of the secondary wall, which will be called "Slignification", proceeds mainly after the development of a secondary wall framework, that is, in the final (F) stage of differentiation, although its initiation can be detected already during the S2 stage.

> To obtain more detailed information, the immunogold-labeling technique has been applied to differentiate between macromolecular features of condensed (mainly C–C bonds) and noncondensed (*O*-4 aryl–alkyl bonds) lignin subunits during deposition. The results obtained with

a young poplar tree suggest that condensed G and GS types of lignin substructures are preferably formed in middle lamella and cell corners of the differentiating fibers, whereas the formation of non-condensed bonds is favorable in the incipient S1, just at the onset of secon‐ darization [36]. Recently, a polyclonal antibody against a specific condensed lignin substruc‐ ture, that is, the 8-ring dibenzodioxocin, has been raised for TEM-immunogold detection of the lignification process in cell walls of softwood xylem [33, 34, 37]. The results demonstrated the absence of the dibenzodioxocin structure in very young tracheids where secondary cell wall layers were not yet formed. Moreover, the dibenzodioxocin structure was more abundant in the secondary cell wall layers than in the middle lamella during secondary cell wall thickening. In contrast to the concept of late lignification, that is, in which wood tracheid cell walls undergo lignification only after deposition of polysaccharides, this is an indication that lignification possibly occurs parallel to polysaccharide deposition. In addition to direct lignin labeling, key enzymes involved in the lignification of plant cell walls were localized in developing walls with specific antibodies. In developing xylem cells of Populus, strong labeling of peroxidases in cell corner regions during early developing stages is probably confirming the onset of lignification in these wall portions [38].

### **3.2. Lignin distribution**

### *3.2.1. Distribution of lignin in softwoods*

A number of topochemical detections established that the compound middle lamella is more highly lignified than the secondary wall in typical softwood tracheids [15, 17, 19, 20, 39]. Moreover, the SEM-EDXA technique provided quantitative information of lignin distribu‐ tion with relatively high accuracy. The distribution of lignin in loblolly pine (*Pinus taeda* L.) tracheids was determined by bromination coupled with SEM-EDXA. It is interesting to note that the lignin concentration in the S2 layer is lower than that in either the S1 or S3 layer [40]. Similar finding was also reported for tracheids of Japanese fir *(Abies suchalinen‐ sis* Fr. Schm.) [41].

#### *3.2.2. Distribution of lignin in hardwoods*

In contrast to the tracheid as the main cell in softwoods, hardwoods have a variety of cells, such as vessels, parenchyma, and fibers. The lignin distribution between secondary wall and middle lamella in hardwood fibers is similar to that in softwoods; however the secondary wall of hardwood fibers is often less lignified than the secondary wall of softwood tracheids. Figure 7 shows the distribution of lignin in *populous nigra* stem as determined by TEM with potassium permanganate staining [5]. The TEM image exhibits the inhomogeneous distribution of lignin.

#### *3.2.3. Distribution of lignin in reaction woods*

Reaction woods appear on leaning stems or branches by any force such as a landslide or snowfall. In softwoods, the reaction wood forms at the lower side of leaning stems or branches, where the compression stress reacts on the xylem. Therefore, this reaction wood is generally called compression wood. Compression wood differs from normal wood in its anatomical

Ultrastructure and Topochemistry of Plant Cell Wall by Transmission Electron Microscopy http://dx.doi.org/10.5772/60752 295

a young poplar tree suggest that condensed G and GS types of lignin substructures are preferably formed in middle lamella and cell corners of the differentiating fibers, whereas the formation of non-condensed bonds is favorable in the incipient S1, just at the onset of secon‐ darization [36]. Recently, a polyclonal antibody against a specific condensed lignin substruc‐ ture, that is, the 8-ring dibenzodioxocin, has been raised for TEM-immunogold detection of the lignification process in cell walls of softwood xylem [33, 34, 37]. The results demonstrated the absence of the dibenzodioxocin structure in very young tracheids where secondary cell wall layers were not yet formed. Moreover, the dibenzodioxocin structure was more abundant in the secondary cell wall layers than in the middle lamella during secondary cell wall thickening. In contrast to the concept of late lignification, that is, in which wood tracheid cell walls undergo lignification only after deposition of polysaccharides, this is an indication that lignification possibly occurs parallel to polysaccharide deposition. In addition to direct lignin labeling, key enzymes involved in the lignification of plant cell walls were localized in developing walls with specific antibodies. In developing xylem cells of Populus, strong labeling of peroxidases in cell corner regions during early developing stages is probably

A number of topochemical detections established that the compound middle lamella is more highly lignified than the secondary wall in typical softwood tracheids [15, 17, 19, 20, 39]. Moreover, the SEM-EDXA technique provided quantitative information of lignin distribu‐ tion with relatively high accuracy. The distribution of lignin in loblolly pine (*Pinus taeda* L.) tracheids was determined by bromination coupled with SEM-EDXA. It is interesting to note that the lignin concentration in the S2 layer is lower than that in either the S1 or S3 layer [40]. Similar finding was also reported for tracheids of Japanese fir *(Abies suchalinen‐*

In contrast to the tracheid as the main cell in softwoods, hardwoods have a variety of cells, such as vessels, parenchyma, and fibers. The lignin distribution between secondary wall and middle lamella in hardwood fibers is similar to that in softwoods; however the secondary wall of hardwood fibers is often less lignified than the secondary wall of softwood tracheids. Figure 7 shows the distribution of lignin in *populous nigra* stem as determined by TEM with potassium permanganate staining [5]. The TEM image exhibits the inhomogeneous distribution of lignin.

Reaction woods appear on leaning stems or branches by any force such as a landslide or snowfall. In softwoods, the reaction wood forms at the lower side of leaning stems or branches, where the compression stress reacts on the xylem. Therefore, this reaction wood is generally called compression wood. Compression wood differs from normal wood in its anatomical

confirming the onset of lignification in these wall portions [38].

**3.2. Lignin distribution**

*sis* Fr. Schm.) [41].

*3.2.1. Distribution of lignin in softwoods*

294 The Transmission Electron Microscope – Theory and Applications

*3.2.2. Distribution of lignin in hardwoods*

*3.2.3. Distribution of lignin in reaction woods*

**Figure 7.** TEM images showing the inhomogeneous distribution of lignin, taken at 80 kV. (a) The CC and P showed higher electron density than the adjacent S1. (b) The outer and inner parts of the S2 layer appear more electron dense (stipplings) than the mid part, the density being particularly pronounced in the curved region of the wall (arrows). (c) The lignin distribution in the S2 layer is distinctly inhomogeneous, with the wall appearing to be a mosaic of electrondense (arrowheads) and electron-lucent (arrows) regions. The lucent regions have a pattern of sinuous features along the radial directions. (d) The dark staining of the vessel indicated that it is highly lignified, V: vessel, F: fiber.

appearance. Differences include a more lignified secondary cell wall (S2L) layer, absence of an S3 layer, and the presence of intercellular spaces in the cell corner region [42, 43]. The distri‐ bution of lignin in compression wood has been extensively investigated. Compression wood shows marked changes in the distribution of lignin across the cell wall with reduced lignifi‐ cation of the middle lamella and increased lignification of the S2L layer. In mild compression wood, the lignification of the CCML and the S2L regions is generally comparable, while the S1 and S2 layers were less lignified (Fig. 3) [20]. In severe compression wood, intercellular spaces reduce the contribution of middle lamella lignin to overall lignin content, which is nevertheless increased by the greater lignification of the S2L layer.

On the contrary, reaction wood named as tension wood is formed at the upper side of leaning stems or branches in hardwoods where the xylem loads the tensile stress. Ten‐ sion wood is characterized by the presence of a cellulose abundant gelatinous layer (GL) forming part of the secondary wall in fibers [44-46]. In maple and oak TW fibers, the GL was divided into concentric sub-layers that appeared either as single rings or as several concentric zones of high and low contrast. Weak staining with potassium permanganate was also visualized at the interface of adjoining concentric layers in maple but was more widespread across the GL in oak, indicating the deposition of aromatic compounds within the cellulose structure of the GL [47].

### **4. Hemicelluloses deposition**

Hemicelluloses are a heterogeneous group of polysaccharides, including xyloglucans, xylans, mannans, and glucomannans. They form physical and chemical bonds to cellulose and lignin and therefore have an important role in building the three-dimensional structures of plant cell walls [48]. The detailed structure of the hemicelluloses and their abundance vary widely among species and cell types. Combination of TEM and immunolabeling has provided detailed information about the deposition of main hemicelluloses related to tissue development and differentiation.

#### **4.1. Hemicelluloses deposition in softwoods**

Glucomannans (GMs) are the most abundant hemicelluloses found in softwoods. GMs are composed of a linear backbone of randomly β-(1,4)-linked D-glucosyl and D-mannosyl residues. The ratio of glucosyl and mannosyl units in softwood GMs is approximately 1:3, and D-galactosyl residues are occasionally attached to the backbone with α-(1,6)-glycosidic bonds. In addition to the galactosyl side chain of GMs, softwood GMs also contain partially substi‐ tuted hydroxyl groups with *O*-acetyl groups at C-2 and C-3 of the mannosyl residues [49].

Many studies have reported the distribution of GMs in the softwood cell wall using various immunochemical probes specific to GMs in combination with TEM. Using the enzyme-gold complex method, Joseleau and Ruel (1984) have demonstrated that GMs of spruce (*Picea abies*) are present mainly in secondary walls but not in the compound middle lamella [50]. The distribution of GMs in the differentiating tracheid cell wall of *Chamaecyparis obtuse* was also investigated by immunogold labeling [51]. The electron microscopic observation showed that labeling of GMs was restricted to the secondary walls of the tracheids and the labeling density temporarily increased and then decreased in the outer and middle layers of the secondary wall during cell wall formation. Investigation of Lodgepole pine (*Pinus contorta var. latifolia*

 Englem.) differentiating secondary xylem showed GM deposition not only in the secondary cell wall, but also in the Golgi apparatus, including vesicles [52]. Recently, the detailed spatial and temporal distribution of GMs in differentiating tracheid cell walls of *Cryptomeria japoni‐ ca* was investigated using immunogold labeling in conjunction with TEM, and the influence of acetylation on immunolocalization of GMs was reported (Fig. 8) [53]. At the primary cellwall formation stage in tracheids, GM labeling was absent in the cell wall. GMs began to deposit at the corner of the cell wall in the early S1 formation stage. Compared to the stronger GM labeling present in the innermost and outermost parts of the S1 layer, the middle of the S1 layer showed only minimal labeling, and then increased gradually during S1 formation. Thus, the authors speculate that some softwood GMs may show an intussusceptional deposition mode by penetrating into the intermicrofibrillar spaces during cell wall formation without binding to microfibrils. A clear uneven distribution of GMs in the S2 layer during S2 formation was also observed. GM labeling increased gradually in the S2 layer during S2 formation with the innermost part of the S2 layer showing the highest density of GM labeling. The deposition of GMs in S3 layer also showed a similar trend with weak labeling at the early stage of S3 formation, and then increased labeling during maturation. The increased GM labeling during cell wall maturation is not consistent with the previous study of the differentiating tracheid cell wall of *Chamaecyparis obtuse* suggesting that the density of GM labeling decreased gradu‐ ally during cell wall formation because of lignin deposition during cell wall maturation. GM labeling in mature tracheids of *Cryptomeria japonica* showed higher concentration of GMs in the S1 layer than that in the S2 layer, which is in agreement with immunogold labeling study of *Chamaecyparis obtusa*. In addition, GM labeling was also observed in the CML of *Cryptomeria japonica* mature tracheids whereas GMs showing absence in the CML in either developing or mature tracheid cell walls of *Chamaecyparis obtuse* [51, 53]. To explore the influence of acetyla‐ tion on immunolocalization of GMs, specimens treated with mild alkali solution were also investigated. Deacetylation of GMs with mild alkali treatment led to a significant increase in GM labeling, suggesting that some GM epitopes may be masked by acetylation. It is interesting to note that the changes in GM labeling after deacetylation were not very pronounced until the early stages of S2 formation, indicating that GMs deposited in the cell wall at early stages of cell wall formation may contain fewer acetyl groups than those deposited at later stages. Furthermore, the decreased GM labeling in mature tracheids suggests that some acetyl groups may be removed from GMs after cell wall formation [53].

cation of the middle lamella and increased lignification of the S2L layer. In mild compression wood, the lignification of the CCML and the S2L regions is generally comparable, while the S1 and S2 layers were less lignified (Fig. 3) [20]. In severe compression wood, intercellular spaces reduce the contribution of middle lamella lignin to overall lignin content, which is

On the contrary, reaction wood named as tension wood is formed at the upper side of leaning stems or branches in hardwoods where the xylem loads the tensile stress. Ten‐ sion wood is characterized by the presence of a cellulose abundant gelatinous layer (GL) forming part of the secondary wall in fibers [44-46]. In maple and oak TW fibers, the GL was divided into concentric sub-layers that appeared either as single rings or as several concentric zones of high and low contrast. Weak staining with potassium permanganate was also visualized at the interface of adjoining concentric layers in maple but was more widespread across the GL in oak, indicating the deposition of aromatic compounds within

Hemicelluloses are a heterogeneous group of polysaccharides, including xyloglucans, xylans, mannans, and glucomannans. They form physical and chemical bonds to cellulose and lignin and therefore have an important role in building the three-dimensional structures of plant cell walls [48]. The detailed structure of the hemicelluloses and their abundance vary widely among species and cell types. Combination of TEM and immunolabeling has provided detailed information about the deposition of main hemicelluloses related to tissue development and

Glucomannans (GMs) are the most abundant hemicelluloses found in softwoods. GMs are composed of a linear backbone of randomly β-(1,4)-linked D-glucosyl and D-mannosyl residues. The ratio of glucosyl and mannosyl units in softwood GMs is approximately 1:3, and D-galactosyl residues are occasionally attached to the backbone with α-(1,6)-glycosidic bonds. In addition to the galactosyl side chain of GMs, softwood GMs also contain partially substi‐ tuted hydroxyl groups with *O*-acetyl groups at C-2 and C-3 of the mannosyl residues [49].

Many studies have reported the distribution of GMs in the softwood cell wall using various immunochemical probes specific to GMs in combination with TEM. Using the enzyme-gold complex method, Joseleau and Ruel (1984) have demonstrated that GMs of spruce (*Picea abies*) are present mainly in secondary walls but not in the compound middle lamella [50]. The distribution of GMs in the differentiating tracheid cell wall of *Chamaecyparis obtuse* was also investigated by immunogold labeling [51]. The electron microscopic observation showed that labeling of GMs was restricted to the secondary walls of the tracheids and the labeling density temporarily increased and then decreased in the outer and middle layers of the secondary wall during cell wall formation. Investigation of Lodgepole pine (*Pinus contorta var. latifolia*

nevertheless increased by the greater lignification of the S2L layer.

the cellulose structure of the GL [47].

296 The Transmission Electron Microscope – Theory and Applications

**4. Hemicelluloses deposition**

**4.1. Hemicelluloses deposition in softwoods**

differentiation.

In addition to GMs, the distribution of xylans in tracheid walls was also investigated. Xylans in wood cell walls are basically composed of a backbone of xylose units that are linked by β- (1-4)-glycosidic bonds. Softwood xylans that are called arabino-4-*O*-methylglucuronoxylans (AGXs) contain arabinofuranose units linked by α-(1-3)-glycosidic bonds to the xylan back‐ bone [49]. The distribution of AGXs in differentiating earlywood tracheid cell walls of *Cryptomeria japonica* was systematically investigated using immune-TEM [54]. Xylans were found to first deposit in the corner of the S1 layer in the early stages of S1 formation in tracheids. In addition, large amount of xylans were also observed in CCML from the early stage of cell wall formation. During S1 formation, the innermost S1 layer showed weaker xylan labeling than did the rest of the cell wall. A similar pattern was observed during secondary cell wall formation, with the innermost layer and the boundary between the S1 and S2 layers showing

**Figure 8.** Immunogold localization of glucomannans (GMs) in differentiating and differentiated tracheids. (a) Expan‐ sion cells. (b) Tracheids at the early S1 formation stage. (c) Tracheids at the S1 layer formation stage. (d) Tracheids at the early S2 formation stage. (e) Tracheids at the formation of the S2 layer. (f) Tracheids at the formation of the S3 lay‐ er. (g, h) Mature tracheids.

weaker labeling than other parts of the cell wall. These results indicate the spatial consistency between xylan deposition and lignin deposition in the early stages of tracheid cell wall formation [55]. However, an almost uniform distribution of xylans throughout the entire cell wall was observed in mature tracheids.

Furthermore, to extend the understanding of distributional diversities of hemicelluloses among cells, the deposition of GMs and AGXs in ray cells and pits was investigated by immunolabeling [56]. In comparison with tracheids, ray cells have different deposition processes of GMs and AGXs. GM labeling in ray cells began to be detected at the early stage of S1 formation in tracheids, whereas AGX labeling began to be detected in ray cells at the S2 formation stage in tracheids. In mature ray cells, GM labeling was absent in the innermost layer of ray cells, whereas AGXs were uniformly distributed in the entire ray cell walls. In pits, GM labeling was detected in pit membranes at an early stage of pit formation, but disappeared during pit maturation, indicating that enzymes capable of GM degradation may be involved in pit formation. In contrast to GM labeling, AGX labeling was not observed in pit membranes during the entire pit developmental process.

#### **4.2. Hemicelluloses deposition in hardwoods**

weaker labeling than other parts of the cell wall. These results indicate the spatial consistency between xylan deposition and lignin deposition in the early stages of tracheid cell wall

er. (g, h) Mature tracheids.

298 The Transmission Electron Microscope – Theory and Applications

**Figure 8.** Immunogold localization of glucomannans (GMs) in differentiating and differentiated tracheids. (a) Expan‐ sion cells. (b) Tracheids at the early S1 formation stage. (c) Tracheids at the S1 layer formation stage. (d) Tracheids at the early S2 formation stage. (e) Tracheids at the formation of the S2 layer. (f) Tracheids at the formation of the S3 lay‐

In hardwoods, *O*-acetyl-4-*O*-methylglucuronoxylans (AcGXs) are the main hemicellulose, occupying about 30% of total cell wall components. AcGXs consist of a β-(1,4)-xylan backbone, decorated with acetyl groups and side chains of 4-*O*-methyl-α-D glucuronic acid [49]. The localization of xylans has been studied by various in situ labeling methods, for example, the xylanase-gold method [57, 58], xylanase and anti-xylanase antibodies [59], and the immuno‐ gold method [60-62]. In differentiating xylem of Japanese beech, xylan deposition started in the middle of the S1 layer formation stage and labeling of GXs was seen only in the secondary walls of xylem cells, but not in the primary walls or the middle lamella. In addition, the increased labeling density during cell wall formation strongly suggested that the deposition of GXs may occur in a penetrative way [60]. The distribution of xylans in differentiating *Populus* xylem cells has been systematically investigated using immuno-microscopic methods in combination with monoclonal antibodies (LM10 and LM11) specific to β-(1-4)-linked xylopyr‐ anosyl residues [62]. LM10 antibody binds low-substituted xylans (lsAcGXs), whereas LM11 antibody binds high-substituted xylans (hsAcGXs) in addition to lsAcGXs [63]. Xylan depo‐ sition was detected earliest in fibers at the cell corner of the S1 layer, and then later in vessels and ray cells, respectively. During secondary cell wall development of fibers, xylan deposition began from the cell corner of the S1 layer after initiation of S1 formation and different labeling patterns of LM10 and LM11 antibodies were observed. LM10 showed stronger xylan locali‐ zation in the outer secondary cell wall than inner layer, while LM11 showed uniform xylan labeling in the whole secondary cell wall. Differentiating vessels showed similar patterns of xylan labeling as fibers except that vessels showed more uniform labeling in the mature cell wall with stronger labeling of lsAcGXs than fibers. In ray cells, xylan labeling occurred at the S2 formation stage in fibers, which was much later than that in fibers and vessels, but was also detected at the beginning of secondary cell wall formation in ray cells. Unlike fibers and vessels, ray cells showed a more homogeneous composition and distribution of xylans than fibers and vessels. All of the three pit types in the secondary xylem of aspen (including fiber–fiber, vessel– vessel, and ray–vessel) showed strong labeling of hsAcGXs during differentiation, and yet gradually disappeared during pit maturation.

#### **4.3. Hemicelluloses deposition in gramineous species**

Most of the existing research about distribution of hemicelluloses in gramineous species concentrate on Arabidopsis, which is one of the most frequently used model plants in plant science. Several immunocytochemical studies have reported the distribution of xylans in Arabidopsis stem [64-66]. Xylan deposition in xylary fibers (fibers) was initiated at the cell corner of the S1 layer and the xylan labeling increased gradually during fiber maturation. Metaxylem vessels showed more developed stages of secondary cell wall formation than fibers, but revealed almost identical xylan labeling patterns to fibers during maturation. The consistency of the immunolabeling patterns between LM10 and LM11 in the cell wall of fibers, vessels, and protoxylem vessels indicated that vascular bundle cells may be chemically composed of a highly homogeneous xylan type. In contrast, interfascicular fibers showed different labeling patterns between the two antibodies and also between different develop‐ mental stages. Immunolocalization studies of mannans in Arabidopsis stems have shown that mannans are distributed in the various cell types with different concentrations [67-69]. Temporal and spatial variations in mannan labeling between cell types in the secondary xylem of Arabidopsis stems were examined using immunolocalization with mannan-specific monoclonal antibodies (LM21 and LM22). Mannan labeling in secondary xylem cells (except for protoxylem vessels) was initially detected in the cell wall during S2 formation and increased gradually during development. Labeling in metaxylem vessels (vessels) was detected earlier than that in xylary fibers (fibers), but was much weaker than fibers. The S1 layer of vessels and fibers showed much less labeling than the S2 layer. Some strong labeling was also detected in pit membranes of vessel pits.

### **5. Conclusions**

The potential of TEM for investigation of plant cell walls has already been demonstrated on various plant tissues. The high spatial resolution allows detection of changes in the ultrastruc‐ ture and cell wall polymer deposition on the cell and cell wall level. Nevertheless, complex sample preparation procedure will limit its extensive application, especially in living plant tissues. Thus, when combined with other in situ microscopic techniques (such as atom force microscopy, confocal laser microscopy, confocal Raman microscopy), much more information hidden in plant cell wall will be illustrated.

### **Author details**

Xia Zhou, Dayong Ding, Jing Ma, Zhe Ji, Xun Zhang and Feng Xu\*

\*Address all correspondence to: xfx315@bjfu.edu.cn

Beijing Key Laboratory of Lignocellulosic Chemistry/MOE Key Laboratory of Wooden Ma‐ terial Science and Application, Beijing Forestry University, Beijing, China

### **References**

**4.3. Hemicelluloses deposition in gramineous species**

300 The Transmission Electron Microscope – Theory and Applications

pit membranes of vessel pits.

hidden in plant cell wall will be illustrated.

Xia Zhou, Dayong Ding, Jing Ma, Zhe Ji, Xun Zhang and Feng Xu\*

terial Science and Application, Beijing Forestry University, Beijing, China

\*Address all correspondence to: xfx315@bjfu.edu.cn

**5. Conclusions**

**Author details**

Most of the existing research about distribution of hemicelluloses in gramineous species concentrate on Arabidopsis, which is one of the most frequently used model plants in plant science. Several immunocytochemical studies have reported the distribution of xylans in Arabidopsis stem [64-66]. Xylan deposition in xylary fibers (fibers) was initiated at the cell corner of the S1 layer and the xylan labeling increased gradually during fiber maturation. Metaxylem vessels showed more developed stages of secondary cell wall formation than fibers, but revealed almost identical xylan labeling patterns to fibers during maturation. The consistency of the immunolabeling patterns between LM10 and LM11 in the cell wall of fibers, vessels, and protoxylem vessels indicated that vascular bundle cells may be chemically composed of a highly homogeneous xylan type. In contrast, interfascicular fibers showed different labeling patterns between the two antibodies and also between different develop‐ mental stages. Immunolocalization studies of mannans in Arabidopsis stems have shown that mannans are distributed in the various cell types with different concentrations [67-69]. Temporal and spatial variations in mannan labeling between cell types in the secondary xylem of Arabidopsis stems were examined using immunolocalization with mannan-specific monoclonal antibodies (LM21 and LM22). Mannan labeling in secondary xylem cells (except for protoxylem vessels) was initially detected in the cell wall during S2 formation and increased gradually during development. Labeling in metaxylem vessels (vessels) was detected earlier than that in xylary fibers (fibers), but was much weaker than fibers. The S1 layer of vessels and fibers showed much less labeling than the S2 layer. Some strong labeling was also detected in

The potential of TEM for investigation of plant cell walls has already been demonstrated on various plant tissues. The high spatial resolution allows detection of changes in the ultrastruc‐ ture and cell wall polymer deposition on the cell and cell wall level. Nevertheless, complex sample preparation procedure will limit its extensive application, especially in living plant tissues. Thus, when combined with other in situ microscopic techniques (such as atom force microscopy, confocal laser microscopy, confocal Raman microscopy), much more information

Beijing Key Laboratory of Lignocellulosic Chemistry/MOE Key Laboratory of Wooden Ma‐


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**Ultrastructural and Morphological Description of the Three Major Groups of Freshwater Zooplankton (Rotifera, Cladocera, and Copepoda) from the State of Aguascalientes, Mexico**

Marcelo Silva-Briano, Araceli Adabache-Ortiz, Gerardo Guerrero-Jiménez, Roberto Rico-Martínez and Guadalupe Zavala-Padilla

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/60659

#### **Abstract**

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306 The Transmission Electron Microscope – Theory and Applications

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An ultrastructural and morphological description of the three major groups of freshwater zooplankton (Rotifera, Cladocera, and Copepoda) from the state of Aguascalientes using scanning electron microscopy (SEM) and transmission electron microscopy (TEM) was performed. The main characteristics used for identification keys for each group were particularly investigated and also the cellular morphology of rods and spermatozoids in males of the rotifer *Brachionus bidentatus* has also been investigated. It is noteworthy to mention that in the state of Aguascalientes, three endemic species of rotifers new to science have been described: *Keratella mexicana, Brachionus araceliae*, and *Brachionus josefinae*. Regarding the suborder Cladocera, the analysis of the first and second pair of antenna, rostrum, cephalic pores, postabdomen, and the five pairs of swimming legs has resulted in the description of seven species new to science from the state of Aguascalientes: four species of *Macrothrix,* two species of *Alona*, and one species of *Karualona*. Regarding the subclass Copepoda, four species of Cyclopoida group new to science have been described from Aguascalientes. The taxonomical description of these species included the morphological analysis of the buccal parts and the five pairs of swimming legs with emphasis on the fifth pair of

© 2015 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

legs. The ultrastructural and morphological analysis of each characteristic has been an exhaustive task. The use of SEM and TEM was crucial to identify all these new species. SEM has allowed focusing in the study of new micro-details that have been used for taxonomical clarity, while TEM allows for studies of cellular composition and the physiological functioning of these zooplankton species. The state of Aguas‐ calientes inventory today comprehends more than 100 rotifer species and about 50 cladoceran and 30 copepod species (of which 14 were new to science in all three groups), leading us to believe that the number of species for this inventory could be increased, adding new species to science, in the process.

**Keywords:** Rotifera, Cladocera, Copepoda, Scanning electron microscopy (SEM), Taxonomy, Transmission electron microscopy (TEM)

### **1. Introduction**

Electronic microscopy (EM) has different elements for its applications; all of them have produced important contributions to the different branches of science. While transmission electron microscopy (TEM) has produced an enormous advance in the study of the different cellular components as well as their function, the scanning electron microscopy (SEM) has helped in the recognition of surfaces of cells, tissues, and structures, developing a new way of more detailed study [1].

In the case of limnology, the use of SEM has been of great importance for taxonomists as a tool that allows for a more detailed study of the different planktonic organisms that are mainly microscopic, and thus optical microscopy might not allow for a clear distinction of structures of taxonomical importance. In the other hand, nuclear ultrastructure in animals, plants, fungi, and Protoctista was studied by TEM to consider variations of RNP particles that may be related to the initial evolution of posttranscriptional processing [2].

In the state of Aguascalientes, Mexico, with the help of SEM, the ultrastructural study of the taxonomic groups Rotifera, Cladocera, and Copepoda has led to the discovery of species new to science, as well as the observation of new structures previously unnoticed. Fourteen new species of these three taxonomic groups were described in Aguascalientes (Table 1). In spite of the discoveries obtained so far, there are still many water reservoirs that have not been analyzed before which opens the probability of increasing the number of new species records for the State or even species new to science.

#### **1.1. Sample preparation**

To prepare any organism for SEM is an art. To calculate the concentrations of each substance to apply, to measure the exposure time for each substance on the desired organism, and to know the order in which to proceed through this methodology is an alchemic process. For


**Table 1.** Fourteen species new to science described from Aguascalientes, México.

rotifers, cladocerans, and copepods, we have used methodologies previously designed. However, each organism requires slight adequations in this methodology to obtain better results for its observation. In these cases, we have used the next two methodologies.

#### *1.1.1. Preparation of Samples for SEM*

legs. The ultrastructural and morphological analysis of each characteristic has been an exhaustive task. The use of SEM and TEM was crucial to identify all these new species. SEM has allowed focusing in the study of new micro-details that have been used for taxonomical clarity, while TEM allows for studies of cellular composition and the physiological functioning of these zooplankton species. The state of Aguas‐ calientes inventory today comprehends more than 100 rotifer species and about 50 cladoceran and 30 copepod species (of which 14 were new to science in all three groups), leading us to believe that the number of species for this inventory could be

**Keywords:** Rotifera, Cladocera, Copepoda, Scanning electron microscopy (SEM),

Electronic microscopy (EM) has different elements for its applications; all of them have produced important contributions to the different branches of science. While transmission electron microscopy (TEM) has produced an enormous advance in the study of the different cellular components as well as their function, the scanning electron microscopy (SEM) has helped in the recognition of surfaces of cells, tissues, and structures, developing a new way of

In the case of limnology, the use of SEM has been of great importance for taxonomists as a tool that allows for a more detailed study of the different planktonic organisms that are mainly microscopic, and thus optical microscopy might not allow for a clear distinction of structures of taxonomical importance. In the other hand, nuclear ultrastructure in animals, plants, fungi, and Protoctista was studied by TEM to consider variations of RNP particles that may be related

In the state of Aguascalientes, Mexico, with the help of SEM, the ultrastructural study of the taxonomic groups Rotifera, Cladocera, and Copepoda has led to the discovery of species new to science, as well as the observation of new structures previously unnoticed. Fourteen new species of these three taxonomic groups were described in Aguascalientes (Table 1). In spite of the discoveries obtained so far, there are still many water reservoirs that have not been analyzed before which opens the probability of increasing the number of new species records

To prepare any organism for SEM is an art. To calculate the concentrations of each substance to apply, to measure the exposure time for each substance on the desired organism, and to know the order in which to proceed through this methodology is an alchemic process. For

increased, adding new species to science, in the process.

308 The Transmission Electron Microscope – Theory and Applications

Taxonomy, Transmission electron microscopy (TEM)

to the initial evolution of posttranscriptional processing [2].

for the State or even species new to science.

**1. Introduction**

more detailed study [1].

**1.1. Sample preparation**

Specimens were fixed in 4 % formaldehyde, dehydrated in graded series of ethanol, taken to critical point, mounted in an aluminum stub (1 cm high and 1.2 cm in diameter) and covered with gold. To study the trophi in rotifers, organisms of every species were prepared according to the protocol of [3] with slight modifications. Briefly, this protocol consisted of isolating ten females of rotifers in a Petri dish and then adding a drop of sodium hypochlorite and waiting until the lorica was dissolved. Then the females were washed three times with distilled water and mounted in a SEM cylinder. The specimens were observed in a JEOL 5900 LV scanning electronic microscope.

#### *1.1.2. Preparation of Male Specimens for TEM*

We cultured rotifers until we obtained 500 males. They were fixed in 2 % glutaraldehyde (GTA) and 4 % paraformaldehyde (PFA) with 0.16 M phosphate buffer (PBS). Then, they were postfixated with 1 % osmium tetroxide (OsO4). Later, the males were embedded in epoxy resin (EPON) and observed in a JEOL 1010 transmission microscope operated at 80 kv.

### **2. Rotifera**

#### **2.1. General information**

Rotifers are a primary group of small invertebrates ranging from 53 μm to 2 mm that play an important role in freshwater ecosystems; they can also colonize marine and terrestrial ecosystems; they can even be found in plants (Bromeliaceae), mosses, and lichens [4]. They are recognized by three main characteristics: 1) the corona, which is a complex of cilia in the anterior part of the organism and allows the production of water currents that help the animal to capture food; 2) the trophi which is the chewing apparatus made of chitin [5], and its function is to grind food that has been captured; and 3) the foot which is found in the posterior part of the organism and its function is to secrete a sticky substance which allows the animal to adhere to a substratum. However, in some species, the foot can be absent, as in the case of the species of the genus *Asplanchna*.

This phylum is composed of three main classes: The Monogononta with 1,570 species, the Bdelloidea with 461 species, and the Seisonidae with a few marine species [6]. In Mexico, 300 species have been reported [7, 8], and for the state of Aguascalientes, there are 96 species belonging to 33 different genera (Biodiversidad de Aguascalientes 2009). In the state of Aguascalientes, the taxonomic studies so far has focused in the class Monogononta, specifically in the genera *Brachionus* and *Keratella*. These genera are very common and well represented in the state of Aguascalientes as well as worldwide; in both genera, cryptic species or species complexes have been described. Therefore, ultrastructural analysis represents an important tool to elucidate the relationship among species of these two genera. The emphasis in the use of Monogononta is related to the ease in its manipulation for ultrastructural analysis compared with Bdelloidea, where only a few specialists worldwide are able to correctly identify them. However, there are many other genera of rotifers that might be worth to study and to detail ultrastructural differences among cryptic species. That might be the case of the numerous genus *Lecane*.

#### **2.2. Representative characteristics for identification**

Rotifers are divided into loricated and illoricated, which means that the loricated rotifers possess a cover or carapace that protects them, while in the illoricated ones, the tegument is exposed to the medium. Due to this, the identification of rotifers is performed in two different ways:


However, for a complete analysis in certain species, it is necessary to observe the male and the ornamentation of the cysts; and in cryptic species (like the *Brachionus calyciflorus*, *Brachionus plicatilis*, and *Keratella cochlearis* species complexes), a genetic analysis of coxI sequences [10] or data on cross-mating behavior tests [11] is necessary.

Ultrastructural and Morphological Description of the Three Major Groups of Freshwater Zooplankton… http://dx.doi.org/10.5772/60659 311

recognized by three main characteristics: 1) the corona, which is a complex of cilia in the anterior part of the organism and allows the production of water currents that help the animal to capture food; 2) the trophi which is the chewing apparatus made of chitin [5], and its function is to grind food that has been captured; and 3) the foot which is found in the posterior part of the organism and its function is to secrete a sticky substance which allows the animal to adhere to a substratum. However, in some species, the foot can be absent, as in the case of the species

This phylum is composed of three main classes: The Monogononta with 1,570 species, the Bdelloidea with 461 species, and the Seisonidae with a few marine species [6]. In Mexico, 300 species have been reported [7, 8], and for the state of Aguascalientes, there are 96 species belonging to 33 different genera (Biodiversidad de Aguascalientes 2009). In the state of Aguascalientes, the taxonomic studies so far has focused in the class Monogononta, specifically in the genera *Brachionus* and *Keratella*. These genera are very common and well represented in the state of Aguascalientes as well as worldwide; in both genera, cryptic species or species complexes have been described. Therefore, ultrastructural analysis represents an important tool to elucidate the relationship among species of these two genera. The emphasis in the use of Monogononta is related to the ease in its manipulation for ultrastructural analysis compared with Bdelloidea, where only a few specialists worldwide are able to correctly identify them. However, there are many other genera of rotifers that might be worth to study and to detail ultrastructural differences among cryptic species. That might be the case of the numerous

Rotifers are divided into loricated and illoricated, which means that the loricated rotifers possess a cover or carapace that protects them, while in the illoricated ones, the tegument is exposed to the medium. Due to this, the identification of rotifers is performed in two different

**1.** For the study of illoricated organisms which is the case of the class Bdelloidea or the genus *Asplanchna*, it is necessary to locate the trophi (Figure 1) as it is the best structure to aid in the identification. However, we must record all the characteristics like body shape and count the number of ovaries, vitellarium, and other features in a live uncontracted animal

**2.** For the loricated rotifers, it is necessary to fix them in 4 % formaldehyde. Identification is commonly based on the number and position of the spines as in the genus *Brachionus*; on the shape of the dorsal and ventral valves of the lorica, the different folds, and the shape and structure of the foot and the presence or absence of claws as in the genus *Lecane*; and on the structure and number of the facets of the lorica and the number and disposition of

However, for a complete analysis in certain species, it is necessary to observe the male and the ornamentation of the cysts; and in cryptic species (like the *Brachionus calyciflorus*, *Brachionus plicatilis*, and *Keratella cochlearis* species complexes), a genetic analysis of coxI sequences [10]

of the genus *Asplanchna*.

310 The Transmission Electron Microscope – Theory and Applications

genus *Lecane*.

ways:

**2.2. Representative characteristics for identification**

previous to the fixation of the specimen.

spines in the genus *Keratella* (Figure 1).

or data on cross-mating behavior tests [11] is necessary.

**Figure 1.** Several characteristics used for rotifer identification. The corona and anterior spines of *Brachionus* (A), anteri‐ or spines of *Keratella* (B), corona of *Hexarthra* (C), ramate trophi of a Bdelloid (D), malleate trophi of *Lecane* (E), malleate trophi of *Keratella* (F); malleoramate trophi of *Filinia* (G), incudate trophi of *Asplanchna* (H), malleate trophi of *Brachio‐ nus* (I), the loricated rotifer *Lepadella* with arrow showing pseudosegments of the foot (J), the illoricated rotifer *Hexar‐ thra* showing the arms and setae needed for identification (K), and the loricated rotifer *Keratella* with arrows pointing to the facets of the lorica and anterior and posterior spines needed for identification (L).

#### **2.3. Sexual behavior and morphology of germ line of cells in rotifers**

The variations in the taxonomic characteristics, as well as the peculiar way in which mating behavior occurs in the different genera, have aroused great interest about the knowledge of sexual reproductive behavior in rotifers. Usually class Monogononta reproduces mostly via

**Figure 2.** Monogononta life cycle: When the female produces an amictic egg, then an amictic (parthenogenetic) female is born and this asexual reproduction goes on until some environmental factors trigger sexual reproduction. However, when a mictic (sexual) female is born, it produces a mictic egg that can be fertilized to produce a resting egg (cyst), or if the egg is not fertilized, this unfertilized mictic egg produces a male. The presence of males in the environment allows cross-mating that results in production of sexual eggs or cysts (that are still known as resting eggs). Cysts represent a strategy to overcome harsh environmental conditions.

parthenogenesis, but during environmental conditions that still remain controversial, males appear in the environment and sexual reproduction takes place (Figure 2). However, the class Bdelloidea lacks sex completely [12].

The study of sexual reproductive behavior of rotifers has helped taxonomy to clarify the position of some species thought to be cryptic as is the case of *B. araceliae* which some authors suggested that it might belong to *B. bidentatus* [13]. However, it was recognized as a new species after cultivation in the laboratory for several generations showed that it was not a morphotype of *B. bidentatus*, but rather a new species [14]. Later, significant differences between the male of both species were noted [15].

Similar studies on cross-mating tests of species within the same family but belonging to different genera have been reported [11]. Unusual sexual reproductive behaviors and the peculiar life of male rotifers led to the study of the two types of sexual cells present in males (spermatozoa and rods) (Figure 3).

Ultrastructural and Morphological Description of the Three Major Groups of Freshwater Zooplankton… http://dx.doi.org/10.5772/60659 313

**Figure 3.** Cavity of the cilia in the male of *B. bidentatus* surrounded by cytoplasmatic material produced by rods (A) and (B), spermatozoon (C), different views of transversal cuts of spermatozoa (D), rod segregating cytoplasmatic mate‐ rial (E), and transversal cut of a rod (F).

parthenogenesis, but during environmental conditions that still remain controversial, males appear in the environment and sexual reproduction takes place (Figure 2). However, the class

**Figure 2.** Monogononta life cycle: When the female produces an amictic egg, then an amictic (parthenogenetic) female is born and this asexual reproduction goes on until some environmental factors trigger sexual reproduction. However, when a mictic (sexual) female is born, it produces a mictic egg that can be fertilized to produce a resting egg (cyst), or if the egg is not fertilized, this unfertilized mictic egg produces a male. The presence of males in the environment allows cross-mating that results in production of sexual eggs or cysts (that are still known as resting eggs). Cysts represent a

The study of sexual reproductive behavior of rotifers has helped taxonomy to clarify the position of some species thought to be cryptic as is the case of *B. araceliae* which some authors suggested that it might belong to *B. bidentatus* [13]. However, it was recognized as a new species after cultivation in the laboratory for several generations showed that it was not a morphotype of *B. bidentatus*, but rather a new species [14]. Later, significant differences between the male

Similar studies on cross-mating tests of species within the same family but belonging to different genera have been reported [11]. Unusual sexual reproductive behaviors and the peculiar life of male rotifers led to the study of the two types of sexual cells present in males

Bdelloidea lacks sex completely [12].

strategy to overcome harsh environmental conditions.

312 The Transmission Electron Microscope – Theory and Applications

of both species were noted [15].

(spermatozoa and rods) (Figure 3).

It has been observed that these two cells are intimately related with the fertilization process. Some photographs have evidenced that the cytoplasmic substance that secretes the rods adheres to the necks of the spermatozoa. These results are still not clear, but apparently the chamber in which the spermatozoa are found is surrounded by rods waiting for the sperma‐ tozoa to appear. According to [16], rods assist in the sperm delivery during fecundation and are the first element that is introduced in the female. We are working to determine the role of rods in the reproductive mechanism of rotifers using ultrastructural studies. However, we do not have a very conclusive result yet, but it would be an important contribution to the knowledge of sexual reproductive embryology for invertebrates.

**Figure 4.** Three new species of rotifers recorded in Aguascalientes, Mexico. *Brachionus araceliae* dorsal view A, ventral view B*,* anterior spines C, posterior wing D, trophi dorsal view E, ventral view F [14]); *B. josefinae* ventral view G*,* foot H, dorsal spines I, ventral spines J, trophi dorsal view K, ventral view L [18, 19]; *Keratella mexicana* dorsal view M, ven‐ tral view N, plaque O, trophi dorsal view P, and ventral view Q [20, 21].

#### **2.4. New species of rotifers from Aguascalientes, Mexico**

tozoa to appear. According to [16], rods assist in the sperm delivery during fecundation and are the first element that is introduced in the female. We are working to determine the role of rods in the reproductive mechanism of rotifers using ultrastructural studies. However, we do not have a very conclusive result yet, but it would be an important contribution to the

**Figure 4.** Three new species of rotifers recorded in Aguascalientes, Mexico. *Brachionus araceliae* dorsal view A, ventral view B*,* anterior spines C, posterior wing D, trophi dorsal view E, ventral view F [14]); *B. josefinae* ventral view G*,* foot H, dorsal spines I, ventral spines J, trophi dorsal view K, ventral view L [18, 19]; *Keratella mexicana* dorsal view M, ven‐

tral view N, plaque O, trophi dorsal view P, and ventral view Q [20, 21].

knowledge of sexual reproductive embryology for invertebrates.

314 The Transmission Electron Microscope – Theory and Applications

The study of morphology and ultrastructure of rotifers along with studies of sexual repro‐ ductive behavior has led to the description of males new to science as in *Platyias quadricornis* [17] and more importantly, to the description of three species new to science. That is the case of *Brachionus araceliae* [14], *Brachionus josefinae* [18, 19], and *Keratella mexicana* [20, 21] (Figure 4).

**Figure 5.** Several characteristics important for cladoceran identification. The first pair of antennas (A); cephalic pores (B); caudal ornamentation of carapace (C); second pair of antennas and shape of carapace (D); postabdomen, endclaw, basal spine, and lateral fasciculla of setae (E); and dorsal keel (F).

### **3. Cladocera**

#### **3.1. General information**

Cladocerans are small-size crustaceans ranging from 0.2 to 18.0 mm in length. Most species are characterized for having a carapace that covers the entire body except for the head [22]. A clearer and more general description of cladocerans is achieved by taking the water flea (genus *Daphnia*) as a model: an organism of this genus has a body not clearly segmented; it has a head and trunk, but the latest one is covered by the carapace in the anterior part. The first part of antennas is small and the second part is big and visible; in the head, there is a compound eye. Cladocerans have 5 to 6 pairs of swimming appendages; the postabdomen is quite character‐ istic of these crustaceans; in the dorsal part, there is an incubation chamber where eggs are deposited.

There are 600 species of cladocerans worldwide [23]. In Mexico, some authors have estimated 150 species [24], and for the state of Aguascalientes, there are 45 species recorded which are distributed in six families [9]. Cladocerans can be found in lakes, ponds, small rivers, and streams, among others. However, some genera are found in musks, lichens, soil, and saline reservoirs.

**Figure 6.** Typical swimming appendages of a cladoceran. First pair (A), second pair (B), third pair (C), fourth pair (D), fifth pair (E), and sixth pair (F). Scale bar equals 100 μm.

#### **3.2. Representative characteristics for identification**

The basic characteristics used for cladoceran identification are a) head with or without a compound eye, b) ocellus, c) cephalic pores, d) the first and second pair of antennas, e) shape and ornamentation of the carapace, f) postabdomen including the endclaw and a number of natatory setae, and g) the five or six pairs of swimming appendages. There are some extra characteristics that might be important tools to help with species identification, but some‐ times to observe these peculiar features, it is necessary to dissect the organism. These peculiar features are many times crucial to achieve the species level. Such are the cases of a) the cephalic pores of *Alona anamariae* (Figure 5B), b) the absent or very rudimentary eyes of the genus *Spinoalona* [25], and c) the keel in the dorsal part of the carapace of *Macrothrix mexicanus* (Figure 5 F), just to mention a few cases. However, it is always necessary to support the identification with the analysis of all structures including the five or six swimming appendages (see Figure 6).

### **3.3. New species of cladocera from Aguascalientes, Mexico**

Nearly 500 water reservoirs have been examined in the state of Aguascalientes, mainly to determine the biodiversity of zooplanktonic groups with special emphasis in rotifers, cladocer‐ ans, and copepods. The study of cladocerans in the state of Aguascalientes has led to the discovery of seven species new to science. The genus *Macrothrix* is especially well represent‐ ed with four species new to science: *Macrothrix agsensis* [26], *M. mexicanus* [27], *M. sierrafriaten‐ sis* [28], and *M. smirnovi* [29] (Figure 7A, D, E, F, & G). The genus *Alona* contributed with two species new to science: *Alona aguascalentensis* (Figure 7C) and *A. anamariae* [30] (Figure 7B). The genus *Karualona* contributed with one species: *K. penuelasi* [13] (Figure 7C), which is truly endemic and only found in a small part of the state. Besides the description of new species, these studies have provided information about the distribution, endemism, and morphological variation of the different morphotypes that a particular species might show. These ultrastruc‐ ture studies with SEM and TEM have strengthened the identification of certain taxa and provided criteria that can be used in the future to resolve taxonomic disputes within the Cladocera group.

### **4. Copepoda**

**3. Cladocera**

deposited.

reservoirs.

**3.1. General information**

316 The Transmission Electron Microscope – Theory and Applications

Cladocerans are small-size crustaceans ranging from 0.2 to 18.0 mm in length. Most species are characterized for having a carapace that covers the entire body except for the head [22]. A clearer and more general description of cladocerans is achieved by taking the water flea (genus *Daphnia*) as a model: an organism of this genus has a body not clearly segmented; it has a head and trunk, but the latest one is covered by the carapace in the anterior part. The first part of antennas is small and the second part is big and visible; in the head, there is a compound eye. Cladocerans have 5 to 6 pairs of swimming appendages; the postabdomen is quite character‐ istic of these crustaceans; in the dorsal part, there is an incubation chamber where eggs are

There are 600 species of cladocerans worldwide [23]. In Mexico, some authors have estimated 150 species [24], and for the state of Aguascalientes, there are 45 species recorded which are distributed in six families [9]. Cladocerans can be found in lakes, ponds, small rivers, and streams, among others. However, some genera are found in musks, lichens, soil, and saline

**Figure 6.** Typical swimming appendages of a cladoceran. First pair (A), second pair (B), third pair (C), fourth pair (D),

The basic characteristics used for cladoceran identification are a) head with or without a compound eye, b) ocellus, c) cephalic pores, d) the first and second pair of antennas, e) shape and ornamentation of the carapace, f) postabdomen including the endclaw and a number of

fifth pair (E), and sixth pair (F). Scale bar equals 100 μm.

**3.2. Representative characteristics for identification**

#### **4.1. General information**

Copepods have several characteristics by which they can be recognized from other inverte‐ brates. They have a cylindrical shape and a segmented body. In the anterior part, there is a pair of antennules and in the ventral part of the body, there are five pairs of swimming appendages; in the posterior part of the body, there is an abdomen or urosome containing caudal branches. These organisms are very diverse, and there are more than 11,500 species worldwide [31]. In Mexico, 100 species have been described in epicontinental waters from the three main orders: Cyclopoidea, Calanoidea, and Harpaticoidea [24]. In the state of Aguasca‐ lientes, 47 species have been recorded; however, only the orders Calanoidea and Cyclopoida have been studied (Dodson & Silva Briano 1996; [33]; CONABIO-IMAE-UAA, 2008; [35]. Copepods are located in oceans, lakes, and ponds, but they have also been found in musks, lichens, dry leaves, and bromeliads [39]; a few species have been recognized as parasites [22]. In Mexico, the species of Calanoidea and in a lesser proportion Cyclopoida have shown high endemic rates [36]. Therefore, ultrastructural studies can be important to describe new species, separate cryptic species within a species complex, and elucidate restricted distribution patterns.

**Figure 7.** Seven species new to science of cladocerans found in Aguascalientes, Mexico. *Macrothrix agsensis* (A) [26], *Alona anamariae* (B) [30], *Karualona penuelasi* (C) [13], *M. smirnovi* (D) [29], *Macrothrix mexicanus* (E) [27], *A. aguascalenten‐ sis* (F) [30], and *M. sierrafriatensis* (G) [28].

#### **4.2. Representative characteristics for identification**

Identifying a copepod requires closed examination of the fifth swimming appendages of the female for cyclopoids and of the male for calanoids. For harpacticoids, it is required to closely examine the maxillipeda for the initial identification. However, the fifth pair of swimming appendages and the maxillipeda are just two of the many features that have to be analyzed. For the populations of calanoids and cyclopoids, we require to analyze a) the first two pairs of antennas, b) the buccal apparatus (mandible, maxilula, maxillae, and the maxillipeda), and c) the urosome (Figure 8). For calanoids, it is necessary to observe the growth in some appen‐ dages in the last segments of the thorax. Other morphological variations that might be important for taxonomic identification include a) pores in the swimming appendages, b) different patterns of setae, and c) modifications in the second pair of antennas in some parasite copepods.

**Figure 8.** Habitus with arrows pointing to the second pair of antennas, a pair of swimming appendages, and urosome (A), caudal part of the urosome (B), segment of the second pair of antennas (C), the first and second pair of antennas (D), caudal part of the urosome (E), fifth pair of swimming appendages (F), and urosome showing the sixth pair of swimming appendages (G).

**Figure 7.** Seven species new to science of cladocerans found in Aguascalientes, Mexico. *Macrothrix agsensis* (A) [26], *Alona anamariae* (B) [30], *Karualona penuelasi* (C) [13], *M. smirnovi* (D) [29], *Macrothrix mexicanus* (E) [27], *A. aguascalenten‐*

Identifying a copepod requires closed examination of the fifth swimming appendages of the female for cyclopoids and of the male for calanoids. For harpacticoids, it is required to closely examine the maxillipeda for the initial identification. However, the fifth pair of swimming appendages and the maxillipeda are just two of the many features that have to be analyzed. For the populations of calanoids and cyclopoids, we require to analyze a) the first two pairs of antennas, b) the buccal apparatus (mandible, maxilula, maxillae, and the maxillipeda), and

*sis* (F) [30], and *M. sierrafriatensis* (G) [28].

**4.2. Representative characteristics for identification**

318 The Transmission Electron Microscope – Theory and Applications

#### **4.3. New species of copepods from Aguascalientes, Mexico**

The study of copepods in the state of Aguascalientes has been less intense than that of cladocerans and rotifers. This is because the state lacks an established researcher with a permanent position in the main academic and research facilities, a condition that exists for the other two taxonomic groups. In spite of that, four species new to science have been found in Aguascalientes. That is the case of *Acanthocyclops dodsoni* [33] (Figure 9), *A. marceloi* [38] (Figure 9), *A. caesariatus* [37] (Figure 9), and *Paracyclops hirsutus* [38].

**Figure 9.** Three new species of copepod recorded in Aguascalientes, Mexico. *Acanthocyclops dodsoni*, dorsal view (A); caudal rami, dorsal view (B); fifth pedigerous and genital doble-somites, ventral view (C) (Mercado- Salas et al. 2006); *A. marceloi*, dorsal view (D); fifth pedigerous, ventral view (E), and urosome, ventral view (F) [37]; *A. cesariatus*, dorsal view (G); fifth pedigerous, ventral view (H); and urosome, ventral view (I) [38]. Scale bar: A, C=250 μm; G=200 μm; B, D, F, H=100 μm; I=50 μm; and E=10 μm.

### **5. Importance of the taxonomic study of rotifers, cladocerans, and copepods**

The primary goal of the taxonomical study of rotifers, cladocerans, and copepods is to know the morphology and to carry on an inventory of all species located in a particular geographic area. The second goal is achieved once species are compared regarding morphological similarities and differences which allows for a better system to identify and classify them correctly. However, all these comparisons at the end allow clarification of questions related to evolution and adaptation when we combine ultrastructural morphological, physiological, and genetic studies.

The observation of small modifications among species of these three taxonomic groups only evidences the specificity that exists between the external environment and the internal function of the organism. The perfect design of each species defines the specific niche of each species in an ecosystem. Each slight modification like apparition of small setae in the fifth leg of a copepod, the changes in the structure of a trophi in rotifers, or the presence of a slightly more elongated endclaw in cladocerans is of the greatest importance since such tiny change can be the difference between a species able to perpetuate itself and the others that go extinct. These observations only corroborate the philosophy started by Darwin in his famous book *On the Origin of the Species*. With this philosophy in mind, then we can estimate the socioeconomic importance that each species implies. The precise knowledge of the species distribution, the kind of ecosystems and niches where we found each species, and the way in which each species interact are a necessity to the economical and sustainable use of each species. For example, if one wishes to use copepods as food for fishes, it would be convenient to know which species grow easily in the region. This knowledge would help to curve maintenance costs and to attain a maximal production of the cultured organisms, in the meantime avoiding producing an ecological misbalance. Another clear example is the use of the cladoceran *Daphnia magna* as model organism for acute toxicological test according to the Mexican Norm NMX-AA087- SCFI-2010, in which sometimes we exposed neonates of this organism in situ to determine the survival rate. However, this species is native of Europe and it is adapted to reservoirs with conditions quite different to the Europeans (like Mexico). The use of nonnative species to evaluate the health of Mexican ecosystems only introduces a bias into the experimental results. If instead we are able to identify and culture native species for each region and use it as a model organisms, we would obtain high-quality results and be more prepared to preserve the environmental health of our reservoirs.

Some other questions of even greater transcendence, as well as improvement in experimental and industrial designs, are the results of basic research as it is the case of the taxonomy aided by ultrastructural studies.

### **Acknowledgements**

**4.3. New species of copepods from Aguascalientes, Mexico**

320 The Transmission Electron Microscope – Theory and Applications

9), *A. caesariatus* [37] (Figure 9), and *Paracyclops hirsutus* [38].

The study of copepods in the state of Aguascalientes has been less intense than that of cladocerans and rotifers. This is because the state lacks an established researcher with a permanent position in the main academic and research facilities, a condition that exists for the other two taxonomic groups. In spite of that, four species new to science have been found in Aguascalientes. That is the case of *Acanthocyclops dodsoni* [33] (Figure 9), *A. marceloi* [38] (Figure

**Figure 9.** Three new species of copepod recorded in Aguascalientes, Mexico. *Acanthocyclops dodsoni*, dorsal view (A); caudal rami, dorsal view (B); fifth pedigerous and genital doble-somites, ventral view (C) (Mercado- Salas et al. 2006); *A. marceloi*, dorsal view (D); fifth pedigerous, ventral view (E), and urosome, ventral view (F) [37]; *A. cesariatus*, dorsal view (G); fifth pedigerous, ventral view (H); and urosome, ventral view (I) [38]. Scale bar: A, C=250 μm; G=200 μm; B,

D, F, H=100 μm; I=50 μm; and E=10 μm.

We are thankful to M. Sc. Reyna Lara Martinez who helped us in the management of the TEM JEOL 1010 at the Faculty of Sciences of the Universidad Nacional Autónoma de México.

### **Author details**

Marcelo Silva-Briano1\*, Araceli Adabache-Ortiz1 , Gerardo Guerrero-Jiménez1 , Roberto Rico-Martínez2 and Guadalupe Zavala-Padilla3

\*Address all correspondence to: msilva@correo.uaa.mx

1 Universidad Autónoma de Aguascalientes Laboratorio de Ecología, Depto. Biología, Avenida Universidad, Aguascalientes,

2 Universidad Autónoma de Aguascalientes, Laboratorio de Toxicología Acuática, Depto. Química, Avenida Universidad, Aguascalientes, México

3 Unidad de Microscopía Electrónica. Instituto de Biotecnología, Universidad Nacional Autónoma de México, Av. Universidad, Cuernavaca, México

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\*Address all correspondence to: msilva@correo.uaa.mx

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2 Universidad Autónoma de Aguascalientes, Laboratorio de Toxicología Acuática, Depto.

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### **Chapter 14**
