Establishment of a Primary Culture from Explants
of the Secondary Phloem of the Carrot Root
This article outlines the step-by-step procedure originally developed by the Steward group for obtaining callus cultures from carrot roots, as illustrated in Fig 3.3 This method is versatile and can typically be modified for use with various tissue types.
The preparation of explants from a carrot root involves several essential pieces of equipment: A sterilized aqua dest is used for washing the tissue, while a jar is designated for surface sterilization of the carrot root After sterilization, the carrot root is placed on a cutting platform to obtain root discs, which are then transferred to a Petri dish for collection Sterilized forceps are utilized to handle the root discs, and a Petri dish with filter paper is used for cutting the explants with a troquar or cork borer Finally, the explants are rinsed in a separate jar, and a needle with a loop at the tip is employed for transferring the explants.
Middle , left Cutting discs from the carrot root, right cutting of explants from the disc Bottom , left
Root disc after cutting the explants, right freshly cut carrot root explants
To prepare carrot root discs, first wrap a cutting platform in aluminum foil or a suitable paper bag, then sterilize it by placing it in a drying oven at 150°C for 4 hours.
• Lids to close apertures in the culture vessels are prepared from aluminum foil by hand, and the vessels are labeled according to the design of the experiment
• Preparation of the nutrient medium follows (see below), and adjustment of the pH of the medium with 0.1N NaOH and 0.1N HCl
To prepare stationary cultures, transfer 15 ml of the nutrient medium into each culture vessel using a pipette or a dispenset Additionally, incorporate solid agar at a concentration of 0.8% for optimal results.
• The culture vessel is closed with aluminum foil caps, and sterilized at 1.1 bar and 120°C for 40 min in an autoclave
To prepare for the explantation of carrot roots, it is essential to sterilize various equipment, including several 9 cm Petri dishes lined with 3–4 layers of filter paper through autoclaving Additionally, one Petri dish should be designated for forceps, a needle, and a troquar, while another dish and two 1-liter beakers require dry sterilization Furthermore, 1 liter of distilled water should be distributed across multiple autoclaved Erlenmeyer flasks For each carrot used in the experiment, two forceps, one troquar, and one platinum or stainless steel needle with a loop, all wrapped in aluminum foil and dry-sterilized, are necessary.
All procedures for acquiring explants for culture are performed in a sterilized inoculation room or preferably on a laminar flow aseptic working bench, which should be activated 30 minutes prior to commencing any experimental work.
To assess the viability and growth potential of carrot explants prior to surface sterilization, a disc matching the diameter of the carrot root is excised and cut using a trochar These explants are then placed in a beaker filled with water; if they float, the root is deemed unsuitable for experimentation In contrast, healthy carrot explants will sink to the bottom of the container, indicating their suitability for further use.
• After the selection of a suitable carrot, the root is scraped and washed with aqua dest., dried with a paper towel, and wrapped into 3–4 layers of paper towel
To sterilize a carrot for processing, place it in a 1-liter beaker and cover it with a sterilizing solution, such as 5% hypochlorite, for 15 minutes It is essential to wear sterile gloves during subsequent steps; if gloves are not available, thoroughly wash hands with ethanol or a clinical disinfectant like Lysafaren frequently throughout the procedure.
• The forceps are dipped into ethanol (96%), flamed, and placed into a sterile Petri dish
• Sterilized water is poured into a sterile Petri dish, ready to receive the explants
To begin the procedure, the cover is carefully removed from the cutting platform and positioned at the center of a sterile working bench A sterile Petri dish with a higher rim is then placed directly beneath the cutting platform, accompanied by sterile forceps for handling.
After removing the carrot from the sterilization solution and washing it with sterilized water, 2-mm discs are carefully cut from the root tip using a cutting platform with precise horizontal strokes This technique is essential to ensure the successful extraction of explants from the carrot root tissue, as any deviation from a horizontal cut may result in contamination from cambium cells instead of the desired secondary phloem.
To obtain the desired number of discs, approximately 15–20 explants can be harvested from the secondary phloem of each disc After this, two forceps are sterilized by flaming and placed into a sterile Petri dish to maintain a contamination-free environment.
To prepare the explants, root discs are carefully transferred using sterile forceps into a Petri dish lined with filter paper Utilizing a sterilized troquar, approximately 20 explants are cut about 2 mm from the cambium These explants are then placed into a Petri dish containing sterilized water It is advisable to cut around 50 additional explants to ensure sufficient quantity for the procedure.
To eliminate any residual sterilizing solution from the roots, explants should be rinsed thoroughly with sterilized water five to six times After the final rinse, most of the water should be removed from the dish, leaving only enough liquid to keep the surface of the explants moist.
To ensure sterile transfer of explants into culture vessels, the needle with a loop at its tip is first dipped in absolute ethanol and then flamed After cooling, the explants are carefully transferred into the nutrient medium without allowing the needle to touch the vessel's opening, preventing microbial contamination Following the transfer to multiple vessels, the needle is flamed again, and the vessels are immediately covered with lids, such as aluminum foil As an additional precaution, both the vessel opening and lid can be flamed before sealing.
Table 3.2 Some disinfectants used in tissue culture experiments, and the concentrations applied (Thorpe and Kamlesh 1984)
• After the work on the laminar flow, the culture vessels with the explants are transferred to the climatized culture room
To obtain sterile cultures from plant material in non-sterile environments, explants can be derived from seedlings grown from sterilized seeds in aseptic conditions Initially, seeds should be immersed in a sterilizing solution for 2–3 hours, preferably using a magnetic stirrer, with the sterilization duration and solution type determined empirically for each plant species Uneven seed coats or hairy coverings may complicate sterilization, but adding a few drops of detergent like Tween 80 can help After surface sterilization, seeds must be rinsed in autoclaved water and then placed on sterilized, moist filters in Petri dishes or sterile agar medium for germination To minimize contamination, it is advisable to limit the number of seeds per vessel When cutting explants from seedlings, a sterilized scalpel or similar device should be used, with frequent flaming to maintain sterility Further procedures involving embryo tissue and immature embryos are discussed in later chapters.
Fermenter Cultures
The principles applied to fermenter or bioreactor cultures can also be utilized in various applications For instance, a bioreactor originally designed for algae cultures has proven effective for culturing cells from several higher plants With the addition of a light source providing approximately 33 W/m², successful investigations into the photosynthesis of photoautotrophic cultured cells in a sugar-free medium have been conducted.
The bioreactor contains 4 liters of nutrient medium that has been sterilized in a vertical autoclave and is placed in the culture room for three days to confirm successful autoclaving before cell transfer If indole-3-acetic acid (IAA) is present in the medium, the fermenter must be kept in the dark to prevent photooxidation After three days, if the bioreactor remains sterile, cell material is transferred using a sterile glass funnel and a 1-cm diameter silicon pipe, ensuring the funnel is positioned in the sterile air stream of the laminar flow in the inoculation cabinet For optimal culture growth, an inoculation of approximately 30 grams of fresh weight is recommended for the 4 liters of medium.
An effective alternative for sterilization involves separately sterilizing the container and the nutrient medium The nutrient medium is sterilized using the previously outlined method, while the empty container undergoes autoclaving for 35 minutes at 1 bar and 130°C For harvesting, the bioreactor contents are poured through layers of fine cheesecloth.
A bioreactor designed for culturing plant material must ensure sufficient mixing while reducing shearing stress and hydrodynamic pressure Since the 1970s, significant advancements have been made in the development of airlift bioreactors, which have proven to be the most effective design for meeting these criteria Notably, the bioreactor has demonstrated minimal damage during the production of somatic embryos from Daucus and Datura cell suspensions used for scopolamine or atropine production.
To reduce operational costs, several disposable plastic devices have been developed as alternatives to reusable glass containers One notable example is the pre-sterilized Life Reactor™ system, created by M Ziff from the Hebrew University and R Levin from Osmototek, a company specializing in advanced plant tissue culture products This system is available in 1.5-liter and 5-liter volumes, with the 1.5-liter vessel capable of producing up to 1,000 plantlets per liter of liquid medium Its user-friendly design makes it ideal for research and small commercial laboratories.
The bioreactor, designed for carrot cell suspension cultivation, operates efficiently within a compact V-shaped bag made from durable plastic laminate, allowing for large-scale multiplication with minimal labor and cost It features a porous bubbler at the bottom for sterile, humidified air supply, and a 1.5-inch inoculation port at the top for adding and withdrawing plant material, sealed with an autoclavable cap The cap includes two ports: one for exhausting excess air and another covered by a silicon rubber septum for aseptic additions.
Immobilized Cell Cultures
In addition to batch cultures, efforts have been made to develop continuous systems in bioreactors, where cells are fixed on a stationary carrier Unlike animal cells, which can autonomously adhere to surfaces like glass or synthetic materials, plant cells face challenges due to their rigid cell walls A potential solution to this issue involves capturing plant cells within the interior of the carrier material.
Calcium-alginate was initially used as a carrier for cell cultivation, but various polymers such as agar, agarose, polyacrylamide, and gelatin have also been tested, along with synthetic materials like polyurethane, nylon, and polyphenyloxide Among these, polyurethane is often favored due to its large inner volume (97% w/v) and the passive invasion mechanism that allows cells to capture within its structure The carrier must be submerged in a cell suspension, enabling cells to divide and grow within the foam's pores until the entire volume is occupied This method does not require additional chemicals for cell fixation, and it has shown no adverse effects on cell vitality and metabolism Polyurethane remains stable in standard nutrient media, even over extended experimental durations, and the cells immobilized on it can be transferred to flatbed containers or columns with a continuous flow of nutrient media.
Circular bioreactor setups that facilitate medium reuse have proven effective for cultivating microbes These continuous systems are designed to produce compounds associated with the primary or secondary metabolism of plant cells, as highlighted by research from Yin et al (2005).
In 2006, research indicated that cell suspensions can be enhanced through immobilized cultures, such as those of Juniperus chinensis, which utilized a 3% alginate gel to effectively produce podophyllotoxin (Premjet and Tachibana 2004) This method allows for modifications in the nutrient medium composition at specific culture stages to optimize cell production Further details will be discussed in Chapter 10.
Nutrient Media
Nutrient media are crucial for the success of cell culture systems, as not all plant organs and tissues can grow autotrophically While most higher plants thrive in light with adequate water and mineral nutrients, roots and developing seeds depend on assimilates and phytohormones from shoot tissues to survive and grow effectively.
Cell culture systems derived from intact plants require a nutrient medium to substitute for substances sourced from various plant parts with unique metabolic functions While some cultures can grow photoautotrophically in inorganic media, most are heterotrophic or mixotrophic, particularly after chloroplast development To meet energy and synthesis needs, carbohydrates such as mono- or disaccharides are typically added, with sucrose being the most common choice However, the effectiveness of carbohydrate supplements varies based on tissue type, research objectives, and plant species, necessitating preliminary investigations to determine the optimal carbohydrate for each specific culture.
Tables 3.3 to 3.7 present the composition of various nutrient media currently in use, highlighting that sucrose concentration typically ranges from 2% to 3% Additionally, the mineral salts mixture, a crucial element of these nutrient media, has evolved significantly since the initial introduction of plant cell nutrient media.
Fig 3.5 Plant cells in poly- urethane foam (photograph by
Table 3.3 Compositions of some nutrient media in use for plant cell and tissue cultures
(for 1 l aqua dest.; the compositions of the stock solutions are given in Table 3.4)
The nutrient media formulations BM, MS, NL, NN, and B5 are essential for plant tissue culture, each containing specific components to support growth For instance, sucrose concentrations vary, with BM, NL, NN, and B5 containing 20 g, while MS has a higher concentration of 30 g Casein hydrolysate is included in NN and B5 at 0.20 g and 0.25 g, respectively, while glycine is present in BM and MS at 1.0 ml All formulations utilize a mineral solution of 100 ml, with varying amounts of Fe and Mg solutions to enhance nutrient availability Vitamins are included in all media, with notable additions of folic acid and biotin in NN and B5 Hormonal supplements such as 2,4-D, IAA, kinetin, and BAP are incorporated in specific media to regulate growth, while m-inositol and coconut milk provide additional nutrients The pH levels of the media are consistently around 5.5 to 5.7, and agar is uniformly added at 8.0 g across all formulations These nutrient media are referenced from various studies, including those by White (1954), Murashige and Skoog (1962), Neumann (1966), Nitsch and Nitsch (1969), and Gamborg et al (1968).
Table 3.4 Compositions of some nutrient media in use for plant cell and tissue cultures: stock solutions
Hormones (mg/100 ml aqua dest.) m-Inositol solution – 500.00 500.00 – 500.00
Table 3.5 Concentrations of some amino acids in casein hydrolysate (as mg/l nutrient medium, by an application of 200 ppm per liter nutrient medium)
Amino acid Concentration Amino acid Concentration
Table 3.6 Concentrations of mineral nutrients in some nutrient media used for cell and tissue culture (final concentration at the beginning of culture, mg/l nutrient medium)
Nutrient medium a BM b MS NL NN B5
In 1954, P White's culture medium was significantly enhanced by the Steward group at Cornell University through the introduction of coconut milk, which increased phosphorus concentration by approximately tenfold This enhancement also led to elevated levels of various mineral components and the addition of micronutrients such as copper and molybdenum While White's medium relied solely on nitrate for inorganic nitrogen, light-grown cultures require 8–10 days to develop functional chloroplasts capable of efficiently reducing nitrite Additionally, glycine served as a source of reduced nitrogen in White's basal medium, but the inclusion of casein offers a more potent alternative.
Table 3.7 Final concentrations of organic components in some media used for plant cell and tissue culture at the beginning of the experiment (mg/l)
Nutrient medium a BM MS NL NN B5
Nutrient medium a BM b MS NL NN B5
Cobalt is present at a concentration of 0.01 in the nutrient medium, which includes 10% coconut milk Various organic nitrogen sources are incorporated, providing 35.4 mg N/l from coconut milk, 0.4 mg from glycine in MS medium, and 31 mg from casein hydrolysate in NL, along with 39 mg in B5 The concentration of hydrolysate in coconut milk is not specified, but it contributes numerous amino acids to the nutrient media Additionally, ammonia is utilized as a source of reduced nitrogen in several media, following the nutrient medium guidelines established by Murashige and Skoog (MS medium).
A key element of nutrient media is a vitamin mixture that typically includes thiamine, pyridoxine, and nicotinic acid Isolated cells from intact plants often lack the ability to produce sufficient amounts of these vital compounds, which are crucial for carbohydrate and nitrogen metabolism However, autotrophic cells can sometimes meet these requirements independently.
Nutrient media, as detailed in Tables 3.3, 3.4, 3.6, and 3.7, include various phytohormones and growth substances that can effectively substitute for coconut milk, which was commonly used in early plant cell cultures The necessity for these phytohormones and their impact on the growth and development of cultured cells largely depends on factors such as the plant species, variety, the type of tissue used for explantation, and the specific objectives of the research Table 3.8 presents examples from our research program demonstrating how different nutrient media with varying hormone supplies can successfully induce primary callus cultures.
Primary explants with meristematic regions can exhibit significant growth in a hormone-free medium, which can be enhanced with auxin application In contrast, explants derived from quiescent tissues, like the secondary phloem of carrot roots, show minimal growth without hormonal stimulation, primarily resulting in cell enlargement The endogenous hormonal systems of cultured explants and their interaction with exogenous hormones from the nutrient medium will be explored in detail in a later chapter, along with the role of hormones in specific cell reactions within various culture systems.
Most nutrient media utilize auxins, primarily naphthylacetic acid (NAA) and 2,4-dichlorophenoxyacetic acid (2,4-D), with indole acetic acid (IAA) sometimes included as a supplement These auxins vary in their chemical and metabolic resistance to breakdown, with IAA being the least stable due to rapid loss from the system through photooxidation and metabolic processes In contrast, NAA offers greater stability, while 2,4-D is the most stable of the three.
In a controlled study over 21 days at 21°C, the growth of explants from the secondary phloem of carrot roots and the pith of tobacco was measured in various liquid nutrient media The original weights of the explants were 3 mg for carrot and 7 mg for tobacco, with results reflecting significant growth in fresh weight per explant across three experimental trials.
Nutrient medium a BM MS NL NN B5
Auxins play a crucial role in stimulating cell division in rapeseed cultures, with IAA treatments promoting the earliest root development For sustained cell division without differentiation, the stable auxin 2.4D is preferred, while IAA or NAA are better for initiating differentiation after a brief cell division phase To prevent adventitious root formation in primary cultures, increasing auxin concentration can be effective However, the optimal use of auxins in nutrient media must be empirically determined for each culture system due to the limited understanding of plant hormonal interactions Additionally, enhancing cell division can be achieved by incorporating cytokinins, such as the synthetic kinetin at low concentrations, while natural cytokinins like zeatin and synthetic options like 6-benzyladenine (BA) may also be utilized in certain culture systems.
M-inositol, a functional component of coconut milk discovered by the Steward group at Cornell University in the early 1960s, is widely used in nutrient media, although its application was first suggested by P White in the early 1950s As a key component of cellular membranes, inositol plays a crucial role in cell signaling systems While it is not classified as a hormone due to its low concentrations in nutrient media, significant hormonal-like responses can still be observed Notably, inositol has been shown to functionally replace indole-3-acetic acid (IAA) under certain conditions, leading to the isolation of IAA–inositol conjugates These conjugates may serve to protect IAA from degradation or inactivate it, similar to the formation of IAA conjugates with glucose.
Table 3.9 presents the effects of various auxins, specifically 2 ppm IAA, 2 ppm NAA, and 0.2 ppm 2,4-D, on the fresh weight, cell count per explant, and rhizogenesis of rapeseed petiole explants (cv Eragi) after 21 days of culture in NL liquid medium, as reported by Elmshọuser in 1977.
0 IAA NAA 2.4D mg F wt./explant 29 51 134 121
Days after beginning of culture – 7 14 42 mg Roots/explant – 13 38 n.d aspartic acid In most cases, m-inositol increases the action of IAA, as well as that of cytokinins
Thermolabile components in nutrient media may be compromised by autoclaving, necessitating the use of sterile filtration for these ingredients In our laboratory, we filter sterilize all components containing radioactive isotopes Additionally, fructose undergoes transformation into toxic substances during autoclaving, which can inhibit the growth of cultured cells, highlighting the importance of filter sterilization.
Evaluation of Experiments
The evaluation of experimental results often involves measuring fresh weight of air-dried material and dry weight after drying at 105°C until a constant weight is achieved To assess the growth of cell suspensions, it is essential to separate the cells and determine the packed cell volume (PCV) A cost-effective and minimally destructive method for this process is utilizing a hand centrifuge at low speeds along with calibrated centrifuge beakers.
To differentiate between growth through cell division and cell enlargement, tissue must be macerated to assess the cell count in a specific tissue sample This is accomplished by counting individual cells within a defined volume of cell suspension using a hemocytometer under a microscope.
To prepare tissue for maceration, it is first frozen at -20°C for 24 hours After thawing, the tissue is immersed overnight in a maceration solution composed of 0.1N HCl and 10% chromic acid in a 1:1 ratio Prior to counting the cells, the macerated tissue is crushed with a glass rod and repeatedly drawn through a syringe.
To count cells effectively, an aliquot of the macerate is placed on a counting grid, with ten counts typically performed per treatment For maceration, a few hundred milligrams of fresh weight are usually sufficient If the maceration process is unsuccessful or yields unsatisfactory results, adjustments to the maceration solution may be necessary based on the specific tissue being studied Notably, many callus cultures and most cell suspensions do not require prior freezing The volume of the maceration solution should be carefully considered to ensure optimal results.
10 times the fresh weight of the tissue to be macerated in mg For calculation:
In the study, VK represents the volume of the grid in microliters, while N indicates the cell number per explant The average cell count, denoted as X, is derived from multiple counts Mv refers to the volume of the maceration solution, and f wt signifies the fresh weight of the macerated tissue measured in milligrams Additionally, n represents the total number of explants that have undergone maceration.
Maintenance of Strains, Cryopreservation
Maintaining certain cell strains viable for extended periods, sometimes up to several years, is often essential for in-depth research that connects various metabolic areas, necessitating the use of a consistent genome.
Regularly setting up subcultures is essential for maintaining cell strains or cell lines, as it addresses the growth of cultures, depletion of nutrient medium components, and the accumulation of waste or dead cells The frequency of subculturing varies based on these factors and can range from a few days to several months or even years.
In Phalaenopsis cultures, which release significant amounts of polyphenols into the medium, subculturing is required every 2 to 3 days In contrast, Arachis cell cultures, which grow slowly through photoautotrophy, only need subculturing every 6 to 8 weeks Typically, these subcultures are maintained stationary on agar.
A widely applicable method for various plant species involves aseptically transferring 10 to 15 pieces of actively growing callus, each approximately 5 mm in diameter, into a 120 ml Erlenmeyer flask filled with 15 ml of nutrient medium This subculturing process typically occurs every four weeks.
The method in question presents a significant drawback due to the cytological and cytogenetical instability of cell material after multiple passages Additionally, it may lead to variations in metabolic processes and genome organization This approach is typically labor-intensive and demands considerable storage space However, by adjusting culture conditions—such as lowering the temperature, limiting nutrients like sugar, and reducing light intensity—storage can be optimized with less frequent subcultures For cold-tolerant species, temperatures can be decreased to nearly 0°C.
Cryopreservation has been adapted for plant cell cultures to address long-term storage challenges, allowing the preservation of cell suspensions and somatic embryos with a revival success rate of up to 80% after thawing Protocols exist for over 100 plant species, enabling somatic embryos and cell suspensions to be stored for several years at the ultra-low temperature of liquid nitrogen (-196°C), where cell division and metabolic activities cease This method significantly reduces maintenance costs and storage space compared to traditional methods that require multiple sub-culturing.
Cryopreservation primarily aims to inhibit ice crystal formation, which can damage cell membranes The effectiveness of this method varies across different plant species and cultures Typically, smaller cells are more amenable to cryopreservation than larger ones, especially those with a diverse range of cytoplasm to vacuole ratios For a comprehensive overview of this technique, refer to Engelmann (1997).
To prepare for cryopreservation, exponentially growing cell cultures are first transferred to a nutrient medium supplemented with 6% mannitol for 3–4 days to osmotically reduce cell water The cryopreservation mixture is prepared at double the concentration of the working solution, consisting of 1M DMSO, 2M glycerin, and 2M sucrose, with a pH of 5.6–5.8, and is filter sterilized Both the cryopreservation mixture and the cell suspension are chilled on ice for 1 hour before being combined and vigorously shaken to create a viscous solution Subsequently, 1 ml of this cell and cryopreservation mixture is placed into sterile polypropylene vials, which are then cooled in a chilling device, gradually lowering the temperature in 1°C intervals to –35°C for 1 hour before storing the vials in liquid nitrogen.
To effectively revitalize cell material, vials must be quickly thawed in warm water at 40°C, followed by an immediate transfer to agar medium Once growth begins, a subsequent transfer to liquid medium is advisable (Seitz et al 1985) The survival rate for Daucus or Digitalis cells preserved using this method ranges from 50% to 75%, while Panax cultures exhibit a lower survival rate.
Evidently, as already mentioned, variations between species exist
During the 1970s and 1980s, methods for preserving differentiated materials like apices and somatic embryos were developed and later modified These modern techniques involve the rapid removal of freezable water followed by quick freezing, leading to vitrification of cellular solutes and the formation of an amorphous glassy structure that prevents ice crystal formation, which can damage cell structure Key variations include encapsulation–dehydration, vitrification, encapsulation–vitrification, pre-growth desiccation, and droplet freezing Encapsulation–dehydration techniques, which stemmed from earlier artificial seed production research, involve pre-growing cells in high sucrose media, desiccating them to about 20% water content, and then freezing them in alginate beads This method has demonstrated high survival rates for various plant species, including temperate crops like apple and grape, as well as tropical plants like sugarcane and cassava Encapsulation–vitrification combines encapsulation in alginate beads with vitrification solution treatment prior to freezing.
An innovative method for preserving potato apices involves using a cryoprotective medium, where dissected apices are pre-cultured in DMSO for a few hours before being frozen in droplets These droplets are then placed on aluminum foil and stored in liquid nitrogen This technique has been successfully implemented across approximately 150 potato varieties, achieving an average recovery rate of 40%.
Successful cryopreservation techniques, particularly for poplar cells, have been extensively reviewed (Tsai and Hubscher 2004), showcasing their application in somatic embryogenesis to preserve clonal germplasm across 23 coniferous tree species (Touchell et al 2002) While slow cooling methods are predominant, vitrification techniques are also utilized In the realm of fruit trees, Reed (2001) highlights the importance of safe storage, which will become increasingly critical as modified genomes are integrated into plant breeding This advancement is particularly relevant for the commercial production of medicinally significant germ lines, especially those producing recombinant proteins (Imani et al 2002; Hellwig et al 2004; Sonderquist and Lee 2008).
Some Physiological, Biochemical,
The carrot root explant system, initially developed by the Steward group at Cornell University using coconut milk for hormones and nutrients, has evolved to utilize a defined mixture of additives such as IAA, m-inositol, and kinetin, replacing the variable composition of coconut milk (Caplin and Steward 1949; Neumann 1966) By incorporating inorganic nutrients from coconut milk into White's original nutrient medium (1954), researchers achieved comparable growth responses in cultured carrot root explants This chemically defined system enables the characterization of component significance for cell division, growth, and differentiation, facilitating the study of the associated physiological and biochemical processes.
Coconut milk, once crucial for tissue culture systems, can still be beneficial in specific scenarios, particularly when other nutrient media fail to promote explant growth; a 10% v/v supplement of coconut milk may yield positive results To prepare coconut milk, the germination openings of the nut are drilled open, and the liquid is filtered through cheesecloth, sterilized by autoclaving, and then deep-frozen until needed In Europe, particularly Germany, the best coconuts for extracting milk are available from late fall to December When cultured carrot root explants are grown in NL3, they experience an initial lag phase of 5–6 days, followed by an exponential growth phase lasting 2–3 weeks, and then transition into a stationary phase During the exponential phase, cell division predominates, leading to a decrease in average cell size compared to the original explants As cell division slows down, the average cell size increases, eventually approaching that of the original explants The duration of these growth phases varies significantly among different carrot root explants, varieties, and species, with notable differences in cell division activity observed despite identical growing conditions, which will be further explored in relation to hormonal influences and environmental factors in subsequent chapters.
The carrot callus, while appearing morphologically unstructured, reveals distinct anatomical layers upon closer inspection Microscopic examination of 2- to 3-week-old callus cultures shows multiple cell layers, with a peripheral layer of larger cells followed by a broad layer of smaller cells Toward the center, a sheath of larger cells is present, often with remnants of the original explant found at the core In older cultures, a noticeable hole may develop Scanning electron microscope images of the cultured explants highlight unique species-specific differences.
The study presents data on the fresh weight, average cell number, and cell weight of callus cultures derived from the secondary phloem of carrot roots over a 24-day period in NL medium The cultures were supplemented with 50 ppm m-inositol, 2 ppm IAA, and 0.1 ppm kinetin, maintained at 21°C under continuous illumination of 4,500 lux Results are expressed in milligrams of fresh weight per explant, average cell count per explant, and nanograms of fresh weight per cell across the culture duration.
Fig 3.7 Section of a carrot callus after 3 weeks of culture
The scanning electromicrographs depict the surface of a callus culture, showcasing Datura innoxia on the left and Arachis hypogea on the right at the top The bottom images feature Daucus carota, with primary explants shown after 6 days of culture on the left and after 21 days on the right (Photograph by A Kumar).
The surface of carrot callus is compact and composed of morphologically similar cells, while Datura and Arachis cultures exhibit a loosely structured surface with deep channels extending through multiple cell layers Notably, Datura culture shows a clear differentiation in cell form The organization of the explant surface plays a crucial role in the formation of cell suspensions, which will be explored in Chapter 4.
Starting from the sixth day, new cells emerge from within the carrot root callus explants, indicating the initiation of cell division beneath the surface Serial sections of the explants reveal small cells actively dividing and forming random clusters known as meristematic nests, which consist of about 100 cells each Within these nests, smaller cells are found at the center, with cell size gradually increasing toward the periphery, suggesting a decrease in cell division activity as cells age The larger cells at the periphery are derived from the outer regions of the meristematic nests, while the central areas exhibit the highest cell division activity, highlighting the dynamic growth patterns within the callus tissue.
Fig 3.9 Section of a carrot root explant cultured for 6 days (NL medium) Note the two meris- tematic nests ( MN ), and cell fragmentation ( CF )
To understand the growth performance of explants and the key factors involved, it's essential to describe the origin and genesis of meristematic nests Approximately 12 to 15 hours post-explantation and culture initiation, cell division begins in 1 to 2 cell layers beneath the cut surface of the explants, particularly at the periphery This process appears to occur randomly, indicating the initiation of cellular activity crucial for explant development.
Fig 3.10 Initiation of cellular fragmentation in cultured carrot root explants (secondary phloem)
The process of cell division initiation, marked by the appearance of phragmosomes and nuclear division, continues for 1-2 days, reducing the distances between the initial division sites and inducing new division centers Concurrently, previously induced cells also undergo division As a result, after 4-5 days of culture, these initial division sites are found at various developmental stages in close proximity, with no histological differences observed related to the timing of their initiation.
The initial cell divisions arise from the septation of large, vacuolated parenchymatic cells in the original tissue, marked by the formation of cytoplasmic threads known as phragmosomes that traverse the central vacuole The nucleus migrates into these threads, leading to the first cell division, which can generate up to eight daughter cells from a single parenchymatic cell This process of cell division initiation is notably different from that observed in cultured petiole explants, which are used to produce adventitious roots or somatic embryos.
A cytophotometric analysis of DNA content in cells at the periphery of explants reveals a peak of 4C cells 20 hours post-culture initiation, indicating a transition into the G2-phase after the S-phase Initially, cells at explantation are primarily in the G1-phase, and the observed 4C-value signifies DNA doubling and the onset of active cell division Twelve hours later, a resurgence of 2C cells indicates a return to G1, marking the completion of the first synchronized cell cycle passage Notably, hormonal influences have not been observed during this phase in hormone-free media; however, explants in media supplemented with IAA or inositol, especially those with added kinetin, exhibit continued cell division This suggests that these phytohormones promote ongoing cell division in response to the wound reaction initiated at explantation, coinciding with ethylene production shortly after culture initiation.
Shortly after the initiation of culture, explants begin synthesizing IAA, followed by 2iP, establishing an endogenous hormonal system that influences further development in conjunction with exogenous phytohormones from the nutrient medium These external hormones sustain cell division, initially triggered by the wound response during explantation In contrast, explants in hormone-free media experience a halt in cell division after two rounds Hormonal treatments can generate meristematic nests, which are absent in hormone-free conditions, and these nests likely foster the development of the necessary endogenous hormonal system for continued growth of the cultured explants.
When interpreting the results of physiological or biochemical investigations in cell populations, such as callus cultures, it's crucial to recognize that the data represent an average of all cells, with significant individual variations These variations, highlighted through cytological and morphological assessments under a microscope, necessitate careful consideration Synchronization techniques can help mitigate these discrepancies, particularly in cell suspensions This situation mirrors the dichotomy in physics between the stability of macro-objects and the complexity of individual cells, a concept referred to as "quantal microbiology" in microbiology Additionally, environmental disturbances during experiments can impact cellular performance; for instance, using a photometer-microscope to measure carotene distribution in carrot root cells is unreliable due to light intensity damaging the carotene This scenario exemplifies the application of Heisenberg's uncertainty principle in biological experimental setups.
Shortly after the culture begins, there is a significant increase in protein synthesis, which is likely associated with the development of cytoplasmic threads traversing the central vacuole, as previously described.
Using 14 C-labeled leucine as a tracer, researchers characterized the distinct protein specimens synthesized during the early stages of callus induction, revealing a hierarchical sequence of protein synthesis Two-dimensional electropherograms of soluble proteins at the onset of culture were stained with Coomassie brilliant blue for visualization, while fluorograms were produced to identify proteins labeled with radioactive leucine that were synthesized during this process.
Fig 3.11 Hypothetic interaction between hormone supply to the nutrient medium, and an endog- enous hormone system of callus cultures e xogenous horm one s
Induction of cell division endogenous pe rpe tua tion of cell division m e ristem a tic
“ne sts” autonom ous horm one system horm one system
Inte ra ction with stable e xogenous horm one s fre sh e xpla nts
Methods to Establish a Cell Suspension
This article provides a practical example of obtaining a cell suspension, similar to the process used for callus cultures Specifically, shoot explants of Datura innoxia were utilized to generate callus, demonstrating a method that can be easily adapted to various other systems.
K.-H Neumann et al discuss the use of plant cell and tissue culture as a crucial tool in biotechnology, focusing on the synthesis of secondary metabolites The article highlights the establishment of callus cultures derived from shoot tissue, where explants from the youngest internode are precisely cut with a sharp scalpel To ensure sterilization, the cut ends are briefly immersed in liquid paraffin, preventing contamination during the surface sterilization process.
Fig 4.1 Histomorphological characterization of suspension cultures of Datura innoxia (Kibler and
Inoculum preparation involves using a 250 µm filtrate and microclusters of approximately 125 µm, alongside histological structures from secondary clusters cultured with kinetin The process requires aseptic handling in a laminar flow bench, where internode segments measuring 1–2 cm are rinsed multiple times with sterilized distilled water Following this, the paraffin cover and epidermis are carefully removed using sterilized scalpels, and the tissue is sliced into 1 mm thick segments These segments, which weigh about 7–8 mg and contain approximately 30,000 cells each, are then halved to establish callus cultures, ensuring a larger size compared to those used for primary carrot cultures.
Fig 4.2 Growth of haploid ( top ) and diploid ( bottom ) cell suspensions of Datura innoxia (Kibler and Neumann 1980) number of cells number of cells in
1n-Suspensionscultures total number of cells secondary calli>250àm number of “free cells” number of “free cells” number of “free cells”