Cell Wall Polysaccharides

The cell wall of angiosperm pollen tubes has been studied extensively (Geitmann 1999; Taylor and Hepler 1997). The highly anisotropic growth behaviour of these cells is reflected in the non-uniform distribution of cell wall components. In the axial direction at least two regions have been identified, the approximately hemispherical growing tip and the cylindrical shank. In the radial direction, depending on the region, one or several cell wall layers can generally be distinguished. If a parallel is drawn with somatic cells, the wall laid down at the tip can be defined as the primary wall, while that formed in mature parts as secondary wall, because that part of the wall has stopped expanding. The temporal sequence of events in somatic cells is, therefore, transformed into a spatial pattern of features in the pollen tube. It is interesting to note that in isolated pollen tube protoplasts the sequence of cell wall deposition events seems to be reversed compared to the original cell. In pollen tubes pectins are laid down first, followed by callose deposition, whereas in isolated pollen tube protoplasts of olive the first cell wall polymer that reappears is callose, while label for pectins is only observed hours later (Majewska-Sawka et al. 2002). Instead of forming separate layers both cell wall components are intermixed in this case.

The tip region of angiosperm pollen tubes is generally characterized by a single wall layer (Lancelle and Hepler 1992). It is known to be mainly composed of newly synthesized pectins (Geitmann et al. 1995). This pectic layer can continue along the entire length of the pollen tube and forms the outer layer of the cell wall in the pollen tube shank, where a secondary wall layer is generally formed adjacent to the plasma membrane, or it can be gradually replaced by this secondary wall as seems to be the case in Arabidopsis thaliana (Derksen et al. 2002). From the outside, this wall has a porous appearance. The thickness of this primary wall is generally around 100 to 300 nm and in the transmission electron microscope (TEM) it appears distinctly fibrillar.

Pectins are galacturonate-rich, acidic polysaccharides, abundant in an-giosperm pollen tubes. In the pollen tube tip region of angiosperms, pectins have a higher degree of methylesterification than in distal regions as shown with fluorescence label using monoclonal antibodies JIM5 and JIM7 (Figs. 1, 2) (Geitmann et al. 1995; Li et al. 1994). This de-esterification is

Fig. 1 Immunofluorescence label of Solanum chacoense pollen tube for pectins with low degree of methyl-esterification using monoclonal antibody JIM5. Label intensity is stronger at the shank of the tube than at the apex (located at the left). Bar = 10 |im. (Micrograph taken by Elodie Parre)
Fig. 2 Immunofluorescence label of Solanum chacoense pollen tube for pectins with high degree of methyl-esterification using monoclonal antibody JIM7. Label intensity is higher at the apex (located at the left). Bar = 10 |im. (Figure reproduced with permission from Parre and Geitmann 2005b)

caused by the activity of pectin methyl esterases secreted at the apex (Li et al. 2002). A distribution of methyl-esterified pectins similar to the one in an-giosperms was observed in gymnosperm pollen tubes (Derksen et al. 1999a). In Pinus, no acidic pectin was found in the pollen tube except for the site where the inner intine of the pollen grain stretches over the pollen tube. Fluctuations in the growth rate cause the density of pectins to change along the longitudinal axis of the tube resulting in the appearance of more or less regularly spaced rings (Fig. 3). It should be noted that these fluctuations in pectin density are not accompanied by overall changes in cell wall thickness (Geit-mann et al. 1995; Pierson et al. 1995), but after EDTA extraction periodic alternations of material with different material properties are revealed in the scanning electron microscope (Derksen, personal communication).

The "secondary" wall layer appears less electron dense and more homogeneous than the fibrillar outer layer (in TEM) even though stratification has been observed in certain cases (Kroh and Knuiman 1982). This inner layer consists mostly of callose as evidenced in the light microscope by staining with decolorized aniline blue. This stain shows that the polymer starts around

Fig. 3 Immunofluorescence label of Nicotiana tabacum pollen tube for pectins with low degree of methyl-esterification using monoclonal antibody JIM5. Label intensity shows regular fluctuations along the longitudinal axis of the pollen tube. The spatial frequency of the fluctuations corresponds to that of the growth oscillations. Bar = 10 |im. (Figure kindly provided by Mauro Cresti)

Fig. 3 Immunofluorescence label of Nicotiana tabacum pollen tube for pectins with low degree of methyl-esterification using monoclonal antibody JIM5. Label intensity shows regular fluctuations along the longitudinal axis of the pollen tube. The spatial frequency of the fluctuations corresponds to that of the growth oscillations. Bar = 10 |im. (Figure kindly provided by Mauro Cresti)

30 |m from the tip in Nicotiana tabacum (Ferguson et al. 1998). In Papaver rhoeas (Geitmann and Parre 2004) and Arabidopsis thaliana (Derksen et al. 2002), the first deposition of callose was observed closer to the tip which might be related to the lower growth rate in these species as discussed below. In petunia it was shown that the thickness of the callose lining increases gradually towards the base (Herrero and Dickinson 1981) whereas in poppy the level of label density did not change distal of 120 |m from the tip (Geitmann and Parre 2004). The use of monoclonal antibodies in TEM showed that callose is confined to the translucent inner layer of the cell wall (Geitmann et al. 1995; Meikle et al. 1991). Under certain circumstances the callose layer can thicken considerably. This is the case, for example, in self-incompatible pollen tubes that have stopped growing (Geitmann et al. 1995). In Nicotiana pollen tubes callose is a (1,3)-^-D-glucan with a few (1,6)-^-linked glycosyl branches (Rae et al. 1985). Unlike cellulose, callose is not organised as crystalline microfibrils, and is often described as amorphous. In addition to the inner wall layer, callose is also the main component of the callosic plugs that are formed at regular intervals in older tubes. These allow the living cytoplasm of the cell to be concentrated at the tip and separated from distal pollen tube regions that eventually degenerate. In Pinus pollen tubes no translucent inner wall layer is visible at the TEM level and no callose plugs are present (Derksen et al. 1999a).

In angiosperm pollen tubes, cellulose is present in exceptionally low amounts. Schlupmann et al. (1994) calculated that in Nicotiana alata pollen tubes that had grown for at least 4 h, the cellulose content was only 10%, whereas 81% of the neutral polysaccharides was callose. The description of the subcellular localization of cellulose varies depending on the method used and species investigated. While many authors suggested that it is located in the outer layer because of the fibrillar appearance in the TEM of the latter (Shivanna and Johri 1985), immunolabel revealed cellulose to colocalise with callose (Fig. 4) (Ferguson et al. 1998). In grasses, on the other hand, it has

Fig. 4 Transmission electron micrograph of a longitudinal section in an older portion of a Nicotiana tabacum pollen tube. Black dots represent double gold-label with both cel-lobiohydrolase (CBHI) (20-nm gold) for cellulose and anti-(1,3)-^-D-glucan monoclonal antibody (30-nm gold) for callose. The colocalisation of label indicates that cellulose is located in the callosic inner wall layer of the pollen tube. (Figure reproduced with permission from Ferguson et al. 1998)

Fig. 4 Transmission electron micrograph of a longitudinal section in an older portion of a Nicotiana tabacum pollen tube. Black dots represent double gold-label with both cel-lobiohydrolase (CBHI) (20-nm gold) for cellulose and anti-(1,3)-^-D-glucan monoclonal antibody (30-nm gold) for callose. The colocalisation of label indicates that cellulose is located in the callosic inner wall layer of the pollen tube. (Figure reproduced with permission from Ferguson et al. 1998)

Fig. 5 Transmission electron micrograph of longitudinal sections of Nicotiana tabacum pollen tube plugs. a, b Labelled with CBHI for cellulose. Cellulose is associated with the periphery of the plug. Bars = 1 |im (a), 0.2 |im (b)

been reported that cellulose forms a third layer between callose and pectins (Heslop-Harrison 1987). In Nicotiana, cellulose was also found to be associated with the periphery of the callosic plugs (Figs. 5, 6) (Ferguson et al. 1998).

Unlike callose, cellulose is a crystalline component of the cell wall and therefore the orientation of microfibrils is potentially important for cell wall architecture. In Petunia pollen tubes, the direction of cellulose microfibrils seems to be random at the tip of the tube whereas in the shank they show a preferential angle of between + 45° and - 45° (Derksen et al. 1999; Sassen 1964) thus causing the appearance of birefringence. Sassen (1964) remarked also that on the inside of the wall close to the tip, cellulose microfibrils might

Fig. 6 Transmission electron micrograph of longitudinal sections of Nicotiana tabacum pollen tube plugs. a, b Labelled with anti-(1,3)-^-D-glucan monoclonal antibody for callose. Callose represents the main component forming the plug. (Figures reproduced with permission from Ferguson et al. 1998). Bars = 1 |im (a), 0.3 |im (b)

Fig. 6 Transmission electron micrograph of longitudinal sections of Nicotiana tabacum pollen tube plugs. a, b Labelled with anti-(1,3)-^-D-glucan monoclonal antibody for callose. Callose represents the main component forming the plug. (Figures reproduced with permission from Ferguson et al. 1998). Bars = 1 |im (a), 0.3 |im (b)

actually be oriented in transverse direction and suggested that this is the direction in which they were laid down. In angiosperm pollen, the tip is generally not or only weakly labelled for cellulose (Derksen et al. 1999; Ferguson et al. 1998), whereas in Pinus pollen tubes the amount of cellulose in the apex is relatively high (Derksen et al. 1999), which could be either a result or a cause of the low growth rate in these cells.

Taken together, the architecture of the pollen tube cell wall shows features typical of plant cells, such as primary and secondary layers. On the other hand it is rather unusual as it contains an unusually high percentage of callose compared to other plant cells. However, Steer and Steer (1989) pointed out that these data need to be interpreted with caution since almost all quantitative analyses were carried out in in vitro growing tubes. Plant cells are known to be able to switch between the production of cellulose and callose, for example upon wounding. It may be that the pollen tube surrounded by artificial liquid medium reacts by synthesizing a less permeable, callosic wall structure to maintain a particular environment. A first indication that this might be the case is the fact that pollen tubes grown in medium stiffened by agarose show lower quantities of callose than those grown in liquid medium (Parre and Geitmann 2005a). Further support comes from the comparison between in vitro and in vivo grown pollen tubes of Arabidopsis which indicates that the growth environment affects the cell wall. In this species the outer, fibrillar cell wall layer is absent from the basal parts of in vitro grown pollen tubes (Derksen et al. 2002), whereas it is present in the in vivo situation (Lennon and Lord 2000). Quantitative chemical and thorough histological analyses that compare in vitro and in vivo grown pollen tubes still remain to be done, however.

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