Cellulose in secondary xylem of Cactaceae: chemical composition and anatomical distribution

doi:10.21203/rs.3.rs-1657136/v1

Cellulose is the main polymer that gives strength to the cell wall and consists of crystalline-microfibrils and amorphous-matrix cellulose located in the primary and secondary cell walls of the plants. In the Cactaceae family, there are studies on the composition of lignin but not on the composition of cellulose. Therefore, the objective of his work is to analyze the chemical composition and distribution of cellulose in the cells of the xylem tissue. Twenty-five species of Cactaceae were collected, dried, and milled. The extractables were removed until obtaining extractive-free wood. Subsequently, cellulose was purified, readings with Fourier transform infrared spectroscopy were made, and the crystallinity indexes were calculated. In addition, for the anatomical analysis of wood, stem sections for each plant were fixed, cut, and stained with safranin O / fast green to observe them under the epifluorescence microscope with three excitation and emission bands. The results showed that crystalline cellulose percentages were different between non-fibrous and fibrous species. In non-fibrous species, the values for crystalline cellulose were above 60%. The fluorescence emission of cellulose in all species was red, the distribution of cellulose in non-fibrous species was in the primary wall; while in the fibrous species, the distribution was in the internal walls and the pits. The high percentages of crystalline cellulose may be related to its distribution in the non-lignified parenchyma and primary walls of tracheary elements with helical or annular thickenings of non-fibrous species, possibly offering structural rigidity and as part of the defense system against pathogens.

Crystalline cellulose

FTIR

fluorescence microscopy

crystallinity indexes

Cactaceae

The cell wall of plants confers rigidity, delimits the cell space, and functions as a physical barrier against pathogens (Srivastava et al. 2017). The main polymers of the cell wall are cellulose, hemicellulose, and lignin, in addition to other compounds such as pectins, structural proteins, and glycoproteins (Bidhendi et al. 2020). The cell wall divides in two: the primary wall with abundant cellulose microfibers, some hemicelluloses, xyloglucans, and pectins (Kuki et al. 2020). The primary wall develops during cell growth and maintains some elasticity during the initial stage. The secondary wall has three layers, S1, S2, and S3. A secondary wall develops once cell growth ends, lignin accumulates mainly, and cellulose occurs in less quantity than other polysaccharides such as hemicelluloses (Harris 2006).

Cellulose is the main component of the cell wall (Srivastava et al. 2017), it is a homopolysaccharide composed of repeated glucose residues and linked by β-(1→4) bonds that generate long and rigid microfibrils (Lampugnani et al. 2018) conformed of 18 cellulose polymers (Kubicki et al. 2018). The presence of cellulose microfibrils in the cell wall confers mechanical and enzymatic resistance (Zhao and Dixon 2014), structural rigidity mainly in the primary wall (Bidhendi et al. 2020), and forms the scaffolding for binding with pectins and hemicelluloses (Lampugnani et al. 2018). Two types of cellulose are characterized by orientation and packaging, the cellulose formed by microfibrils or also called crystalline cellulose (Festucci-Buselli et al. 2007), which is packaged because the microfibrils are linked by hydrogen β(1→4)-D-GLCP bonds, making it more compact, rigid and ordered (Malinovsky et al. 2014). The cellulose matrix has no order (amorphous) because it forms a network where the hemicelluloses, pectins, lignin, and phenolic compounds are inserted and joined (Mnich et al. 2020; Kuki et al. 2020).

The importance of studying the chemical composition of cellulose is due to its use in the paper industry (Festucci-Buselli et al. 2007) and the production of biofuels (Carere et al. 2008). The species with the highest amount of amorphous cellulose (Ciolacu et al. 2011) and the lowest presence of lignin or lignin with the highest accumulation of syringyl monomers (Alves et al. 2011) are those that have potential for use in the industries previously mentioned. On the other hand, cellulose studies also focus on anatomical-structural analysis to identify cell wall interactions with biotic and abiotic factors (Bacete et al. 2018). At this point, different authors analyzed the stressful effects of the environment on cellulose accumulation (Le Gall et al. 2015) and the interaction between cellulose and pathogens (Kesten et al. 2019).

In succulent species, studies are scarce and most focused on the genus Agave a member of the Asparagaceae family (Hidalgo-Reyes et al. 2015). In the Cactaceae family, few studies identify the chemical composition of lignin (Reyes-Rivera et al. 2015; Maceda et al. 2018, 2019, 2021), further, the composition of cellulose is only analyzed in Opuntia ficus-indica due to the potential of the use of cellulosic compounds in the biofuel and paper industry (Yang et al. 2015). However, of the other species of the Opuntioideae, Pereskioideae, and Cactoideae subfamilies, there are no studies on the composition of cellulose and its distribution in the secondary xylem (wood), which allow us to understand the structure and functioning of cellulose in the cell wall of the parenchyma, water conductive and supporting cells. Therefore, the aims were to analyze the chemical composition of cellulose and its distribution in wood. The hypothesis is that there was variation in the chemical composition of cellulose among the different groups of cacti due to the type of wood present.

Chemical analysis of cellulose

Adult and healthy plants of representative species of Cactaceae (Table 1) were collected. The vascular cylinder of all species was isolated and dried for 72 h. After that, 2 grams of each species were weighed, dried, and grounded. Subsequently, to obtain the free-extractive wood, successive extractions were applied with ethanol:benzene (1:2), ethanol 96%, and hot water at 90º C according to the method proposed by Reyes-Rivera and Terrazas (2017).

Table 1 Morpho-anatomical characteristics of the 25 species of Cactaceae

Species	Collection number	Wood type	Stem
Coryphantha clavata (Scheidw.) Backeb.	BV2535	Non-fibrous	Cylindrical
C. cornifera (DC.) Lem.	BV2534	Non-fibrous	Cylindrical
C. delaetiana (Quehl) A. Berger	BV2542	Non-fibrous	Globose
C. delicata L. Bremer	SA1927	Non-fibrous	Cylindrical
C. hintoriorum Dicht & A. Lüthy	BV2539	Non-fibrous	Cylindrical
C. macromeris (Engelmann) Lemaire	BV2600	Non-fibrous	Globose
C. pallida Britton & Rose	SA860	Non-fibrous	Globose
C. pseudoechinus Boed.	BV2543	Non-fibrous	Cylindrical
C. ramillosa Cutak	HSM3775	Non-fibrous	Globose
C. retusa Britton & Rose	SG55	Non-fibrous	Globose
Echinocereus cinerascens (DC.) Lem. subsp. tulensis	SA1744	Non-fibrous	Cylindrical
E. dasyacanthus Engelm.	SA2077	Non-fibrous	Cylindrical
E. pectinatus (Scheidw.) Engelm.	SA1918	Non-fibrous	Cylindrical
E. pentalophus (DC.) Lem.	SA1740	Non-fibrous	Cylindrical
Mammillaria carnea Zucc. Ex Pfeiff.	DA241	Non-fibrous	Cylindrical
M. dixathocentron Backeb. Ex Mottram	CPNL133	Non-fibrous	Cylindrical
M. magnifica Buchenau	UG1411	Non-fibrous	Columnar
M. mystax Mart.	DA238	Non-fibrous	Cylindrical
Neolloydia conoidea (DC.) Britton & Rose	BV2595	Non-fibrous	Cylindrical
Cylindropuntia imbricata (Haw.) F. M. Knuth	TT990	Fibrous	Tree
C. kleiniae (DC.) F. M. Knuth	TT1000	Fibrous	Shrub
C. leptocaulis (DC.) F. M. Knuth	TT994	Fibrous	Shrub
Opuntia stenopetala Lem.	TT997	Fibrous	Shrub
O. stricta (Haw.) Haw.	TT998	Fibrous	Shrub
Leuenbergeria lychnidiflora (DC.) Lodé	TT967	Fibrous	Tree

With the free-extractive wood, cellulose purification was performed by Maceda et al. (2018) procedure based on the Kûshner-Höffer method. 0.5 g of extractive-free wood were weighed and added to 12.5 mL of HNO3/ethanol (1:4 v/v), kept in a reflux system for 1 h, allowed to cool to room temperature, and decanted the sample. Subsequently, 12.5 mL of HNO₃/ethanol were added again and the process of reflux, cooling, and decantation was repeated two more times. In the last process, 12.5 mL of 1% aqueous KOH solution were added, kept under reflux for an additional 30 minutes, and finally filtered through a fine pore Büchner filter. The residue (cellulose) was dried at 60° C for 12 h.

To identify the presence of crystalline and amorphous cellulose, a Fourier Transformed Infrared Spectrometer (FTIR) (Agilent Cary 630 FTIR) was used. A small amount of dry cellulose from each species was taken and readings in the spectrum range of 4000-650 cm-1 (30 scans with a resolution of 4 cm-1, 15 seconds per sample) of each sample were made with the program MicroLab PC (Agilent Technologies, Santa Clara, CA, United States). Baseline correction and averaged spectra were performed with the Resolution Pro FTIR Software program (Agilent Technologies, Santa Clara, CA, United States). Using the crystallinity indexes applied to the FTIR spectra, the presence of crystalline cellulose and amorphous cellulose were calculated in triplicate for each species using the following indexes:

-Proportion of crystallinity (Ciolacu et al. 2011) or also called total crystallinity (TCI) (Poletto et al. 2014): the area under the absorbance curve of 1372 cm-1 (A₁₃₇₂) and 2900 cm-1 (A₂₉₀₀) was measured, consequently, the ratio of the TCI bands (A₁₃₇₂/A₂₉₀₀) was obtained. The result was multiplied by 100 to obtain the percentage of crystalline cellulose: the closer to 100 %, the greater the amount of crystalline cellulose.

-The lateral order index (LOI) (Poletto et al. 2014) or the second proportion of crystallinity (Ciolacu et al. 2011) was calculated with the area under the curve of absorbances 1430 cm-1 (A₁₄₃₀) and 893 cm-1 (A₈₉₃) to finally obtain the ratio LOI (A₁₄₃₀ / A₈₉₃). The result was multiplied by 100 to obtain the percentage that allowed us to identify the relationship between crystalline and amorphous cellulose: if the value is close to 100, there is more ordered crystalline cellulose (Ciolacu et al. 2011).

Epifluorescence microscopy

Epifluorescent and confocal microscopy are precise techniques to identify the distribution of structural components such as lignin, cellulose, phenolic compounds, starch, and proteins (Donaldson 2020). In the particular case of epifluorescence, it allows the observation of thin sections in plant samples where the tissues are anisotropic (Kitin et al. 2020). The method used to make the observations was based on the results obtained by De Micco and Aronne (2007) and Maceda et al. (2019, 2021) with epifluorescence and safranin O / fast green staining. Representative samples of wood from the base of the stem Cactaceae species were obtained, fixed with a solution of formalin-acetic acid-ethanol (10:5:85) (Ruzin 1999), and washed before dehydrating the samples (Kitin et al. 2020). The non-fibrous species samples were embedded with paraffin and cut with a rotatory microtome, and the fibrous species were cut with a sliding microtome (Maceda et al. 2018). All the samples were stained with safranin O/fast green and mounted in synthetic resin according to Loza-Cornejo and Terrazas (1996).

Due to different structural components of the cell wall presenting autofluorescence and fluorescence emission (Donaldson 2020), three different excitation and emission bands were used at the same time (Maceda et al. 2019), this process allows having images with a “true color” (Donaldson 2020). Therefore, to identify the differences in cellulose distribution in the Cactaceae secondary xylem, a wide-field fluorescence microscope (Zeiss Axio Imager Z2) with Apotome 2.0 (Zeiss Apotome.2), an AxioCam MRc 5 (Zeiss), and a microscope metal halide fluorescence light source (Zeiss HXP 120) were used. The multicolor images were obtained with a triple excitation/emission bandwidth: DAPI (blue) with excitation of 365/10 nm and emission of 445/50 nm, FITC (green) with excitation of 470/40 nm, and emission of 525/50 nm, TRITC (red) with excitation of 546/12 nm and emission of 575-640 nm. Each sample was exposed to fluorescence light with low power for a maximum of one minute as proposed Baldacci-Cresp et al. (2020) to avoid overexposure of the samples and cause their photobleaching. Images were obtained with the Zen Blue 2.5 lite (Zeiss, Germany) program, and brightness adjustments were made to the entire image.

Cellulose composition

Figure 1 illustrates the cellulose spectra and Table 2 shows the allocation of the main bands. The absence of the bands 1269 cm^− 1 corresponding to lignin and hemicellulose, 1595 cm^− 1, 1512 cm^− 1, and 1463 cm^− 1 assigned to lignin and 1735 cm^− 1 corresponding to xylenes and hemicelluloses reflect the effectiveness of the extraction and cellulose purification. The total crystallinity index (TCI) showed that most species had values close to one (Table 3) due to the greater presence of crystalline cellulose. However, almost all species of Cylindropuntia and Opuntia (except C. imbricata) and non-fibrous species C. delaetiana, E. dasyacanthus, and M. carnea showed almost similar values between crystalline and amorphous cellulose. The lateral order index (LOI) reflected the cellulose structure; the values were variable in the species of Cactaceae (Table 3). The species with the major disorder in the structure were C. retusa, E. cinerascens, C. kleiniae, and C. leptocaulis, while in the other species the cellulose structure order prevailed (52–93%).

Table 2

Assignment of FTIR absorption bands for the cellulose of Cactaceae species
Wavenumber (cm^− 1₎	Assignments
3000–3600	OH stretching
2900	CH stretching
1430	CH₂ bending (crystalline and amorphous cellulose)
1372	CH bending
1336	OH in-plane bending (amorphous cellulose)
1317	CH₂ vibration (crystalline cellulose)
1163	C-O-C asymmetrical stretching
893	Out-of-plane asymmetrical stretching of cellulose ring
670	C-O-H out-of-plane stretching

The two crystallinity indexes were high in most of the Cactaceae species, except in some fibrous species of Cylindropuntia and Opuntia as well as in non-fibrous species, C. delaetiana, E. dasyacanthus where the TCI was lower (Table 3). The species with the highest crystallinity index varied in the LOI index because some species had a low order, such as three species of Coryphantha, E. cinerascens, and C. imbricata, which had high crystallinity rates, but a greater disorder in cellulose.

Table 3

Crystallinity indexes with standard deviation to determine the crystalline cellulose of the 25 Cactaceae species
Species	TCI (%)	LOI (%)	Species	TCI (%)	LOI (%)
C. clavata	70 ± 0.01	69 ± 0.02	E. pentalophus	63 ± 0.01	70 ± 0.02
C. cornifera	86 ± 0.01	52 ± 0.01	M. carnea	58 ± 0.01	74 ± 0.01
C. delaetiana	50 ± 0.01	56 ± 0.02	M. dixathocentron	75 ± 0.01	81 ± 0.02
C. delicata	76 ± 0.01	62 ± 0.01	M. magnifica	75 ± 0.02	75 ± 0.02
C. hintoriorum	66 ± 0.01	64 ± 0.01	M. mystax	93 ± 0.02	93 ± 0.02
C. macromeris	71 ± 0.01	66 ± 0.01	N. conoidea	69 ± 0.01	72 ± 0.02
C. pallida	62 ± 0.01	59 ± 0.01	C. imbricata	69 ± 0.01	50 ± 0.01
C. pseudoechinus	83 ± 0.02	65 ± 0.01	C. kleiniae	53 ± 0.01	48 ± 0.01
C. ramillosa	80 ± 0.01	53 ± 0.01	C. leptocaulis	58 ± 0.01	43 ± 0.01
C. retusa	79 ± 0.02	45 ± 0.01	O. stenopetala	57 ± 0.01	58 ± 0.01
E. cinerascens	77 ± 0.02	44 ± 0.01	O. stricta	47 ± 0.01	73 ± 0.02
E. dasyacanthus	51 ± 0.02	52 ± 0.01	L. lychnidiflora	80 ± 0.02	83 ± 0.02
E. pectinatus	71 ± 0.02	77 ± 0.02

Cellulose distribution in cells of the secondary xylem

Figures 2–5 show representative species of fibrous (Pereskioideae and Opuntioideae) and non-fibrous (Cactoideae) wood. In bright-field microscopy images, the lignified cell walls were observed in red tones and the non-lignified in blue tones. With fluorescence microscopy, the fibrous species reflected bluish-green to yellow tones; while in the non-fibrous species, the tones of the lignified walls were greenish. The presence of cellulose and other components of the non-lignified walls reflected reddish tones in all species. Cylindropuntia leptocaulis (Fig. 2a, b) had starch within some parenchyma (P) cells, which showed cyan-colored. The distribution of cellulose was different between fibrous species and non-fibrous species.

In longitudinal sections of Cylindropuntia, Opuntia, and L. lychnidiflora, cellulose in the pits of the VEs (Fig. 3a, b, d) was detected. Besides, the presence of cellulose was mainly in the simple pits of the F of Cylindropuntia and L. lychnidiflora. (Fig. 3a–b, d). The unlignified parenchyma of O. stenopetala in longitudinal sections showed the primary walls with the presence of cellulose (Fig. 3c), whereas, in the lignified parenchyma of Cylindropuntia and Leuenbergeria, lignin accumulated mainly in the cell wall and cellulose was exclusively in the pits (Fig. 3a, b, d).

In non-fibrous species (Fig. 4), cellulose fluorescence emission was detected in the primary wall of P, WBT, and VE, unlike in fibrous species, where fluorescence emission was not detected in the primary wall but the S3 layer of the secondary wall adjacent to the lumen. Furthermore, the non-fibrous species showed secondary walls as helical and annular thickenings in VE and WBT, so the primary wall was visible in the longitudinal sections (Fig. 5). In all non-fibrous species, the P was abundant and unlignified, in addition to having a greater accumulation of cellulose (Fig. 4b, d, f, h; Fig. 5b).

The majority of species studied had above 60% of crystalline cellulose and the distribution of cellulose was in unlignified parenchyma, VE, and WBT, both with secondary walls as helical and annular thickenings.

Crystalline cellulose

The low crystallinity indexes (47–58%) obtained for some species of cacti were similar to those reported for most gymnosperms and some angiosperms. The species with crystallinity indexes greater than 60% such as Coryphantha pallida and Echinocereus pentalophus were similar to most angiosperms reported by Agarwal et al. (2013) and to the bark of the cactus Cereus forbesii, which had 82% crystalline cellulose (Orrabalis et al. 2019).

Opuntia ficus-indica is the species with reports of crystalline cellulose in different parts of the plant: cladodes 27.2% (Yang et al. 2015) and 79% (Maceda et al. 2020), spines 33.81-70% (Vignon et al. 2004; Marin-Bustamante et al. 2017), vascular tissue 22–28% (Greco and Maffezzoli 2015), fruit epidermis 38% (Habibi et al. 2009) and 60% in seeds (Habibi et al. 2008). Maceda et al. (2020) reported for Opuntia streptacantha and O. robusta, percentages of crystalline cellulose of 76% and 74%, respectively, percentages similar to those reported in this study for some fibrous species and similar to the values of O. stenopetala and O. stricta.

Cellulose distribution and chemical composition

The tonalities observed in the fluorescence emission for cellulose and lignin correspond to that reported in other similar studies with safranin O / fast green staining technique and the three excitation bands (De Micco and Aronne 2007; Maceda et al. 2019, 2021; Donaldson 2020; Kitin et al. 2020). This technique allowed the detection of differences in the distribution of both structural compounds (cellulose and lignin) between non-fibrous and fibrous species. The safranin O dye allows the detection of lignin autofluorescence and fluorescence in analyzed tissues (De Micco and Aronne 2007; Kitin et al. 2020), with tonalities blue to green (Donaldson 2020; Baldacci-Cresp et al. 2020). In the case of celluloses and hemicelluloses that do not have autofluorescence as lignin (Donaldson 2020), with safranin O /fast green staining, the fluorescence emission of cellulose can be detected at 570–620 nm and its tonalities were reddish similar to the results of Maceda et al. (2019, 2021). However, subsequent studies with immunohistochemistry or specific fluorophores could help confirm the distribution of cellulose (Donaldson 2020; Bidhendi et al. 2020).

When comparing the results with that reported by Maceda et al. (2019) for the primary xylem, cellulose accumulated in the primary walls and lignin in the secondary ones of the helical and annular thickenings of the protoxylem and metaxylem of fibrous species, similar to that observed in non-fibrous adult plants. Only in the metaxylem of Leuenbergeria lychnidiflora, there was a decrease in the accumulation of cellulose in the primary walls and a greater accumulation in the alternate pits, similar to that obtained in the secondary xylem of the fibrous species.

The presence of high percentages of crystalline cellulose in most of the studied cacti, mainly in non-fibrous species, could be related to the abundant unlignified parenchyma and the lower accumulation of lignin in the xylem as seen in Fig. 4 and Fig. 5b. In fibrous species of Cylindropuntia and Opuntia, there was a higher percentage of crystalline cellulose in vascular tissue compared to non-fibrous species, possibly because xylem has a greater accumulation of lignin in the cell walls of the VE, F, and P (Maceda et al. 2018). This high percentage of lignin may contribute as a physical (Hamann 2012) and chemical barrier against the attack of pathogens (Durkovič et al. 2014). Zhao and Dixon (2014)) and Bacete and Hamann (2020) mentioned that the cell wall is a dynamic barrier in conditions of abiotic stress. Therefore, in some cells, the presence of increased lignin accumulation can be observed (Polo et al. 2020), while in others, similar to the gelatinous layer (G), it mainly accumulates cellulose (Festucci-Buselli et al. 2007).

In this type of cell where only the primary wall is present, the accumulation of crystalline cellulose packed by hydrogen chain β-(1→4)-D-Glcp bonds (Malinovsky et al. 2014) makes cellulose hydrophobic (Festucci-Buselli et al. 2007), confers structural support (Malinovsky et al. 2014) and causes a decrease in the efficiency of cellulase enzymes (Thomas et al. 2013) by not presenting sites binding with enzymes, making it difficult to degrade (Rytioja et al. 2014). In contrast, amorphous cellulose is slightly hydrophilic (Thomas et al. 2013), susceptible, and degrades rapidly due to the action of cellulase enzymes (Suchy et al. 2011) and pH changes by pathogens (Kubicek et al. 2014).

Infection with some pathogenic fungi occurs when the hyphae invade the roots and subsequently the vascular tissue (Kesten et al. 2019). In the fibrous species of Cactaceae, the accumulation of lignin works as a physical barrier in VEs (Liu et al. 2018). However, in non-fibrous species with little accumulation of lignin in helical or annular thickenings, the presence of crystalline cellulose in the primary wall of the tracheary elements and the unlignified parenchyma, would function as a physical barrier to prevent the spread of the fungus by reducing the effectiveness of cellulase enzymes (Thomas et al. 2013). In subsequent studies using transmission electron microscopy techniques (Ruel et al. 2012), the presence of crystalline cellulose in the primary wall of the tracheal elements can be analyzed and characterized, which will support and confirm this assertion.

For Leuenbergeria lychnidiflora, whose accumulation of lignin in vascular tissue is 36% (Reyes-Rivera et al. 2015) to 42.52% (Maceda et al. 2018), the percentage of crystalline cellulose is 80% an,d a cellulose arrangement of 83% similar to that observed in non-fibrous species. These results suggest a possible relationship with the sites where this species is distributed, which is found in areas of higher humidity (Edwards and Donoghue 2006) compared to the other fibrous species (Cylindropuntia and Opuntia) analyzed whose distribution is in arid and semi-arid climates (Gómez-Hinostrosa and Hernández 2000). Further analyses would be necessary to determine whether this percentage of crystalline cellulose in L. lychnidiflora prevails in other Leuenbergeria and Pereskia species, early-diversified taxa of Cactaceae as well as other Opuntia species.

The resistance of plants to stressful conditions is energetically expensive, in addition to the constitutive expression of defense mechanisms, such as the accumulation of callose (Srivastava et al. 2017; Lampugnani et al. 2018), pectins (Bacete and Hamann 2020), or secondary metabolites (Meraj et al. 2020). This is not always the best strategy against colonization of pathogens or diseases, because it can restrict physiological processes and have negative impacts on the plant, such as reduced seed production and biomass (Bacete et al. 2018). Therefore, the presence of primary physical barriers, such as lignin (Miedes et al. 2014) and crystalline cellulose (Malinovsky et al. 2014) that inhibit the spread of pathogens, would decrease the expression of these defensive systems energetically expensive (Bacete et al. 2018). The heterogeneity in the composition of cell walls between species reflects the diversity of defensive mechanisms against degrading enzymes that pathogens have developed to break plant cell walls, such as numerous cell wall-degrading enzymes (CWDEs), polygalacturonases, or xylanases (Annis and Goodwin 1997).

In these Cactaceae species, as mentioned before, cellulose would work to confer structural support on the primary wall without losing flexibility (Moura et al. 2010) and thus maintain the cell structure during periods of water stress and rain (Garrett et al. 2010). Furthermore, the increased amount of crystalline cellulose could function as a defense against pathogens (Thomas et al. 2013), by providing resistance to degradation by glycosyl hydrolase enzymes (Malinovsky et al. 2014) and enzymes produced by pathogenic fungi (Kesten et al. 2019). Therefore, that non-fibrous cacti species would accumulate a greater amount of crystalline cellulose as a primary defense in the xylem against possible pathogen attacks. Further analysis will be necessary to confirm the presence of percentages of crystalline cellulose and its possible polymorphisms by X-ray diffraction (French 2014). In addition to analyzing more species of dimorphic and non-fibrous cacti together with other families of plants, that present succulent stems to determine if the high percentages of crystalline cellulose persist due to the abundance of non-lignified parenchyma.

The non-fibrous species presented more than 60% of crystalline cellulose. This distribution was in the primary cell wall of the tracheary elements and the unlignified parenchyma. In fibrous species, the distribution was in the cell wall near the lumen and the simple and alternate pits of vessel elements and fibers. The high percentages of crystalline cellulose could be related to resistance to pathogens.

Acknowledgments

Funding was provided by DGAPA-UNAM postdoctoral fellowship to AM, and by the DGAPA-PAPIIT, UNAM grants IN209012 and IN210115 to TT. Authors thank to Dr. Pedro Mercado Ruaro from Laboratorio de Morfo-Anatomía y Citogenética (LANABIO, UNAM), for allowing us to use the fluorescence microscope; thank to M. C. Rubén San Miguel Chávez for allowing us to use the FTIR, and Julio César Montero-Rojas for artwork.

Author contributions

TT and AM designed the work, AM, MSH, TT performed the experiments and analyzed the data, AM, and TT wrote the manuscript. All the authors read and approved the manuscript.

Conflict of interest Statement

The authors declare that have no competing interests.

Data availability statement

Raw data of FTIR spectra are available in the Figshare repository: https://doi.org/10.6084/m9.figshare.14301257

Agarwal UP, Reiner RR, Ralph SA (2013) Estimation of cellulose crystallinity of lignocelluloses using near-IR FT-Raman spectroscopy and comparison of the Raman and Segal-WAXS methods. J Agric Food Chem 61:103–113. https://doi.org/10.1021/jf304465k
Alves A, Simoes R, Stackpole DJ et al (2011) Determination of the syringyl/guaiacyl ratio of Eucalyptus globulus wood lignin by near infrared-based partial least squares regression models using analytical pyrolysis as the reference method. J Near Infrared Spectrosc 19:343–348. https://doi.org/10.1255/jnirs.946
Annis SL, Goodwin PH (1997) Production and regulation of polygalacturonase isozymes in Canadian isolates of Leptosphaeria maculans differing in virulence. Can J Plant Pathol 19:358–365. https://doi.org/10.1080/07060669709501060
Bacete L, Hamann T (2020) The role of mechanoperception in plant cell wall integrity maintenance. Plants 9:574. https://doi.org/10.3390/plants9050574
Bacete L, Mélida H, Miedes E, Molina A (2018) Plant cell wall-mediated immunity: cell wall changes trigger disease resistance responses. Plant J 93:614–636. https://doi.org/10.1111/tpj.13807
Baldacci-Cresp F, Spriet C, Twyffels L et al (2020) A rapid and quantitative safranin‐based fluorescent microscopy method to evaluate cell wall lignification. Plant J 102:1074–1089. https://doi.org/10.1111/tpj.14675
Bidhendi AJ, Chebli Y, Geitmann A (2020) Fluorescence visualization of cellulose and pectin in the primary plant cell wall. J Microsc 278:164–181. https://doi.org/10.1111/jmi.12895
Carere CR, Sparling R, Cicek N, Levin DB (2008) Third generation biofuels via direct cellulose fermentation. Int J Mol Sci 9:1342–1360. https://doi.org/10.3390/ijms9071342
Ciolacu D, Ciolacu F, Popa VI (2011) Amorphous cellulose-structure and characterization. Cellul Chem Technol 45:13–21
De Micco V, Aronne G (2007) Combined histochemistry and autofluorescence for identifying lignin distribution in cell walls. Biotech Histochem 82:209–216. https://doi.org/10.1080/10520290701713981
Donaldson L (2020) Autofluorescence in plants. Molecules 25:2393. https://doi.org/10.3390/molecules25102393
Durkovič J, Kačík F, Olčák D et al (2014) Host responses and metabolic profiles of wood components in dutch elm hybrids with a contrasting tolerance to dutch elm disease. Ann Bot 114:47–59. https://doi.org/10.1093/aob/mcu076
Edwards EJ, Donoghue MJ (2006) Pereskia and the origin of the cactus life-form. Am Nat 167:777–793
Festucci-Buselli RA, Otoni WC, Joshi CP (2007) Structure, organization, and functions of cellulose synthase complexes in higher plants. Brazilian J Plant Physiol 19:1–13. https://doi.org/10.1590/s1677-04202007000100001
French AD (2014) Idealized powder diffraction patterns for cellulose polymorphs. Cellulose 21:885–896. 10.1007/s10570-013-0030-4
Garrett TY, Huynh C-V, North GB (2010) Root contraction helps protect the “living rock” cactus Ariocarpus fissuratus from lethal high temperatures when growing in rocky soil. Am J Bot 97:1951–1960. https://doi.org/10.3732/ajb.1000286
Gómez-Hinostrosa C, Hernández HM (2000) Diversity, geographical distribution, and conservation of Cactaceae in the Mier y Noriega region, Mexico. Biodivers Conserv 9:403–418. https://doi.org/10.1023/A:1008935710910
Greco A, Maffezzoli A (2015) Rotational molding of biodegradable composites obtained with PLA reinforced by the wooden backbone of Opuntia ficus-indica cladodes. J Appl Polym Sci 132:42447. https://doi.org/10.1002/app.42447
Habibi Y, Heux L, Mahrouz M, Vignon MR (2008) Morphological and structural study of seed pericarp of Opuntia ficus-indica prickly pear fruits. Carbohydr Polym 72:102–112. https://doi.org/10.1016/j.carbpol.2007.07.032
Habibi Y, Mahrouz M, Vignon MR (2009) Microfibrillated cellulose from the peel of prickly pear fruits. Food Chem 115:423–429. https://doi.org/10.1016/j.foodchem.2008.12.034
Hamann T (2012) Plant cell wall integrity maintenance as an essential component of biotic stress response mechanisms. Front Plant Sci 3:77. https://doi.org/10.3389/fpls.2012.00077
Harris PJ (2006) Primary and secondary plant cell walls: a comparative overview. New Zeal J For Sci 36:36–53
Hidalgo-Reyes M, Caballero-Caballero M, Hernández-Gómez LH, Urriolagoitia-Calderón G (2015) Chemical and morphological characterization of Agave angustifolia bagasse fibers. Bot Sci 93:807–817. https://doi.org/10.17129/botsci.250
Kesten C, Gámez-Arjona FM, Menna A et al (2019) Pathogen‐induced pH changes regulate the growth‐defense balance in plants. EMBO J 38:e101822. https://doi.org/10.15252/embj.2019101822
Kitin P, Nakaba S, Hunt CG et al (2020) Direct fluorescence imaging of lignocellulosic and suberized cell walls in roots and stems. AoB Plants 12:1–19. https://doi.org/10.1093/aobpla/plaa032
Kubicek CP, Starr TL, Glass NL (2014) Plant Cell Wall–Degrading enzymes and their secretion in plant-pathogenic fungi. Annu Rev Phytopathol 52:427–451. https://doi.org/10.1146/annurev-phyto-102313-045831
Kubicki JD, Yang H, Sawada D et al (2018) The shape of native plant cellulose microfibrils. Sci Rep 8:13983. https://doi.org/10.1038/s41598-018-32211-w
Kuki H, Yokoyama R, Kuroha T, Nishitani K (2020) Xyloglucan is not essential for the formation and integrity of the cellulose network in the primary cell wall regenerated from Arabidopsis protoplasts. Plants 9:629. https://doi.org/10.3390/plants9050629
Lampugnani ER, Khan GA, Somssich M, Persson S (2018) Building a plant cell wall at a glance. J Cell Sci 131:jcs207373. https://doi.org/10.1242/jcs.207373
Le Gall H, Philippe F, Domon J-M et al (2015) Cell wall metabolism in response to abiotic stress. Plants (Basel Switzerland) 4:112–166. https://doi.org/10.3390/plants4010112
Liu Q, Luo L, Zheng L (2018) Lignins: Biosynthesis and biological functions in plants. Int J Mol Sci 19:335. https://doi.org/10.3390/ijms19020335
Loza-Cornejo S, Terrazas T (1996) Anatomía del tallo y de la raíz de dos especies de Wilcoxia Britton & Rose (Cactaceae) del noreste de México. Bot Sci 59:13–23. https://doi.org/10.17129/botsci.1502
Maceda A, Reyes-Rivera J, Soto-Hernández M, Terrazas T (2021) Distribution and chemical composition of lignin in secondary xylem of Cactaceae. Chem Biodivers 18:e2100431. https://doi.org/10.1002/CBDV.202100431
Maceda A, Soto-Hernández M, Peña-Valdivia CB et al (2019) Differences in the structural chemical composition of the primary xylem of Cactaceae: A topochemical perspective. Front Plant Sci 10. https://doi.org/10.3389/fpls.2019.01497
Maceda A, Soto-Hernández M, Peña-Valdivia CB et al (2020) Characterization of lignocellulose of Opuntia (Cactaceae) species using FTIR spectroscopy: possible candidates for renewable raw material. Biomass Convers Biorefinery. https://doi.org/10.1007/s13399-020-00948-y
Maceda A, Soto-Hernández M, Peña-Valdivia CB, Terrazas T (2018) Chemical composition of cacti wood and comparison with the wood of other taxonomic groups. Chem Biodivers 15:e1700574. https://doi.org/10.1002/cbdv.201700574
Malinovsky FG, Fangel JU, Willats WGT (2014) The role of the cell wall in plant immunity. Front Plant Sci 5:178. https://doi.org/10.3389/fpls.2014.00178
Marin-Bustamante MQ, Chanona-Pérez JJ, Güemes-Vera N et al (2017) Production and characterization of cellulose nanoparticles from nopal waste by means of high impact milling. Procedia Eng 200:428–433. https://doi.org/10.1016/j.proeng.2017.07.060
Meraj TA, Fu J, Raza MA et al (2020) Transcriptional factors regulate plant stress responses through mediating secondary metabolism. Genes (Basel) 11:346. https://doi.org/10.3390/genes11040346
Miedes E, Vanholme R, Boerjan W, Molina A (2014) The role of the secondary cell wall in plant resistance to pathogens. Front Plant Sci 5:358. https://doi.org/10.3389/fpls.2014.00358
Mnich E, Bjarnholt N, Eudes A et al (2020) Phenolic cross-links: building and de-constructing the plant cell wall. Nat Prod Rep. https://doi.org/10.1039/c9np00028c
Moura JCMS, Bonine CAV, de Oliveira Fernandes Viana J et al (2010) Abiotic and biotic stresses and changes in the lignin content and composition in plants. J Integr Plant Biol 52:360–376. https://doi.org/10.1111/j.1744-7909.2010.00892.x
Orrabalis C, Rodríguez D, Pampillo LG et al (2019) Characterization of nanocellulose obtained from Cereus forbesii (a South American cactus). Mater Res 22:20190243. https://doi.org/10.1590/1980-5373-MR-2019-0243
Poletto M, Ornaghi HL, Zattera AJ (2014) Native cellulose: structure, characterization and thermal properties. Mater (Basel Switzerland) 7:6105–6119. https://doi.org/10.3390/ma7096105
Polo CC, Pereira L, Mazzafera P et al (2020) Correlations between lignin content and structural robustness in plants revealed by X-ray ptychography. Sci Rep 10:1–11. https://doi.org/10.1038/s41598-020-63093-6
Reyes-Rivera J, Canché-Escamilla G, Soto-Hernández M, Terrazas T (2015) Wood chemical composition in species of Cactaceae the relationship between lignification and stem morphology. PLoS ONE 10:e0123919. https://doi.org/10.1371/journal.pone.0123919
Reyes-Rivera J, Terrazas T (2017) Lignin analysis by HPLC and FTIR. Methods Mol Biol 1544:193–211. https://doi.org/10.1007/978-1-4939-6722-3_14
Ruel K, Nishiyama Y, Joseleau JP (2012) Crystalline and amorphous cellulose in the secondary walls of Arabidopsis. Plant Sci 193–194:48–61. https://doi.org/10.1016/J.PLANTSCI.2012.05.008
Ruzin SE (1999) Plant microtechnique and microscopy. Oxford University Press, UK
Rytioja J, Hildén K, Yuzon J et al (2014) Plant-polysaccharide-degrading enzymes from Basidiomycetes. Microbiol Mol Biol Rev 78:614–649. https://doi.org/10.1128/mmbr.00035-14
Srivastava V, McKee LS, Bulone V (2017) In: eLS (ed) Plant cell walls. John Wiley & Sons, Ltd, Chichester, UK, pp 1–17
Suchy M, Linder MB, Tammelin T et al (2011) Quantitative assessment of the enzymatic degradation of amorphous cellulose by using a quartz crystal microbalance with dissipation monitoring. Langmuir 27:8819–8828. https://doi.org/10.1021/la2014418
Thomas LH, Trevor Forsyth V, Šturcová A et al (2013) Structure of cellulose microfibrils in primary cell walls from collenchyma. Plant Physiol 161:465–476. https://doi.org/10.1104/pp.112.206359
Vignon MR, Heux L, Malainine M-E, Mahrouz M (2004) Arabinan-cellulose composite in Opuntia ficus-indica prickly pear spines. Carbohydr Res 339:123–131. https://doi.org/10.1016/j.carres.2003.09.023
Yang L, Lu M, Carl S et al (2015) Biomass characterization of Agave and Opuntia as potential biofuel feedstocks. Biomass Bioenergy 76:43–53. https://doi.org/10.1016/j.biombioe.2015.03.004
Zhao Q, Dixon RA (2014) Altering the cell wall and its impact on plant disease: From forage to bioenergy. Annu Rev Phytopathol 52:69–91. https://doi.org/10.1146/annurev-phyto-082712-102237

Cellulose in secondary xylem of Cactaceae: chemical composition and anatomical distribution

Status:

Version 1

Abstract

Figures

Introduction

Material And Methods

Results

Cellulose composition

Cellulose distribution in cells of the secondary xylem

Discussion

Crystalline cellulose

Cellulose distribution and chemical composition

Conclusions

Declarations

References

Status:

Version 1