plasmolysis onion cells experiment

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Plasmolysis: Loss of Turgor and Beyond

Ingeborg lang.

1 Cell Imaging and Ultrastructure Research, University of Vienna, Althanstrasse 14, A-1090 Vienna, Austria; E-Mails: [email protected] (S.S.); [email protected] (B.S.)

Stefan Sassmann

Brigitte schmidt, george komis.

2 CR-Hana, Palacký University Olomouc, Šlechtitelů 586/11, 783 71 Olomouc-Holice, Czech Republic; E-Mail: [email protected]

Associated Data

Plasmolysis is a typical response of plant cells exposed to hyperosmotic stress. The loss of turgor causes the violent detachment of the living protoplast from the cell wall. The plasmolytic process is mainly driven by the vacuole. Plasmolysis is reversible (deplasmolysis) and characteristic to living plant cells. Obviously, dramatic structural changes are required to fulfill a plasmolytic cycle. In the present paper, the fate of cortical microtubules and actin microfilaments is documented throughout a plasmolytic cycle in living cells of green fluorescent protein (GFP) tagged Arabidopsis lines. While the microtubules became wavy and highly bundled during plasmolysis, cortical filamentous actin remained in close vicinity to the plasma membrane lining the sites of concave plasmolysis and adjusting readily to the diminished size of the protoplast. During deplasmolysis, cortical microtubule re-organization progressed slowly and required up to 24 h to complete the restoration of the original pre-plasmolytic pattern. Actin microfilaments, again, recovered faster and organelle movement remained intact throughout the whole process. In summary, the hydrostatic skeleton resulting from the osmotic state of the plant vacuole “overrules” the stabilization by cortical cytoskeletal elements.

1. Introduction

The process of plasmolysis is probably still known to many from their student days. In hyperosmotic solutions such as sucrose, mannitol or sorbitol, water is extruded from the vacuole causing a loss of turgor pressure. If this state persists, the protoplast retracts further, causing the detachment of the plasma membrane from the rigid cell wall. Two major types of plasmolysis are known, depending on: The cell type, the viscosity of the cytoplasm, and the osmoticum used [ 1 ]. In convex plasmolysis, the protoplast is rounded up exhibiting symmetrical convex ends ( Figure 1 a). In concave plasmolysis, the plasma membrane separates from the cell wall by the formation of several concave pockets ( Figure 1 b). Plasmolysis is reversible and the addition of hypotonic solutions or plain water will lead to the re-expansion of the protoplast and the reinstatement of the original turgor pressure [ 1 ]. The central vacuole is the major compartment of osmotic water flow during plasmolysis but obviously, the abrupt change in protoplast size and shape impacts the subcellular architecture as a whole. In this research, we followed the organization of plant cytoskeletal elements namely: Cortical microtubules and actin microfilaments, during a plasmolytic cycle and documented the entire process in living cells.

Schematic of the two major plasmolysis forms; ( a ) convex plasmolysis; ( b ) concave plasmolysis.

Aside from the central vacuole, a rigid cell wall is required for plasmolysis. This structure forms a solid shell encasing the osmo-sensitive and membrane-covered protoplast. Consequently, plasmolysis occurs in walled cell types ranging from bacteria, to fungi and finally plants [ 1 , 2 , 3 , 4 ]. When large molecules with a size above the cell wall exclusion limit are used as osmotica (e.g., polyethylene glycols with a MW above 20 kDa) [ 5 ], then hyperosmolarity induces the collapse of the entire cell wall—plasma membrane continuum in a phenomenon known as cytorrhysis [ 6 ].

In basic biology courses, plasmolysis is used to demonstrate plant cell turgor and its relation to the mechanical rigidity of plant organs. During plasmolysis, the plasma membrane is separated from the cell wall, and this process is easily demonstrated. Specific chemicals like potassium salts lead to swelling of the cytoplasm thereby allowing a distinction between the tonoplast and the plasma membrane in so called cap plasmolysis [ 7 , 8 ]. Furthermore, plasmolysis is an active process characteristic of viable cells; therefore, it is used to test the cellular viability against treatments including heavy metals and other stress factors [ 9 , 10 , 11 ]. The connection of the plasma membrane and the cell wall is still widely discussed subject [ 4 , 12 , 13 , 14 , 15 , 16 ] and plasmolysis is essentially used to analyze the space and link between these two structures.

The term plasmolysis was defined by de Vries [ 17 ] upon the invention of a method to determine a plant’s turgor pressure using hypertonic solutions. Later, Hecht [ 18 ] intensively studied plasmolysed onion epidermal cells. He observed a network-like structure and fine strands (Hechtian strands) in the periplasmic space (the space between the cell wall and the retracted protoplast) that link the protoplast to the inner side of the cell wall of plasmolysed cells. Plasmodesmata have been discussed as candidates for Hechtian attachment sites; these minute channels between adjacent cells have been magnificently observed by transmission electron microscopy in plasmolysed plant tissue [ 19 , 20 ]. However, Hechtian strands can also be formed in the periplasmic space despite cell walls lacking plasmodesmata (e.g., trichomes or outer epidermis walls), suggesting additional structural connections like RGD-containing peptides [ 13 , 14 ], arabinogalactan proteins [ 21 ] or growing cellulose microfibrils [ 22 ]. A comprehensive review on the process of plasmolysis is given by Oparka [ 1 ]. Since then, cytoskeletal elements have been analyzed during a plasmolytic cycle in various plant species and cell types, [ 23 , 24 , 25 ] but mainly in fixed plant material which was used to visualize microtubules and/or actin microfilaments by means of immunolocalization. In this research, the plasmolytic fate of cortical microtubules and actin microfilaments was followed in epidermal hypocotyl cells of appropriate Arabidopsis lines stably transformed with GFP-tagged cytoskeletal markers allowing the documentation of the whole process in vivo .

2. Results and Discussion

2.1. a plasmolytic cycle.

During a plasmolytic cycle, the semipermeable membranes, plasma membrane and tonoplast, were forced to adjust to the loss of water from the vacuole in hypertonic solutions (plasmolysis), or to the water uptake until full turgor is reinstated (deplasmolysis). Plasmolysis started immediately after contact with the plasmolytic solution and in Arabidopsis hypocotyl cells, it was complete after 30 min following exposure to 0.8 M mannitol solution. Cells survived in the plasmolysed state for longer than 24 h, depending on the experimental design and cell type used, [ 26 , 27 , 28 ] while in exceptional cases they recovered after prolonged exposure to hyperosmoticum reinstating turgor and cortical microtubule organization, suggesting the function of volume regulatory increase mechanisms [ 26 ] and references therein, [ 29 ]. Fine Hechtian strands and a network like structure (Hechtian reticulum) provided the contact of the plasma membrane to the cell wall during plasmolysis, while preserving the plasma membrane surplus resulting from protoplast reduction [ 22 , 26 , 30 , 31 ]. The process of plasmolysis was easily observed with good bright field optics. However, the investigation of subcellular changes in living cells required the use of GFP-tagged Arabidopsis lines, as is recognized in the present study using microtubule associated protein 4 (MAP4; [ 32 ]) and tubulin alpha 6 (TUA6; [ 33 ]) lines for labeling microtubules, and a GFP-tagged actin binding domain of fimbrin 1 (ABD; [ 34 ]) for visualization of actin microfilaments in living cells.

2.2. Microtubules

In interphase cells, plasmolysis (which is the disruption of the cell wall—plasma membrane—cortical cytoskeleton continuum) is expected to exert the strongest impact on cortical microtubules since they are closely linked to the plasma membrane, exerting a role in oriented cellulose microfibril deposition [ 15 , 16 , 28 , 35 , 36 , 37 ]. In fully turgid Arabidopsis hypocotyl cells, we observed the biased organization of cortical microtubules arranged in parallel order, and in oblique to transverse orientations in the cortical cytoplasm ( Figure 2 a and Figure 3 f). At the onset of plasmolysis, cortical microtubules became wavy in order to accommodate the decreased shape of the protoplast ( Figure 2 b). Concurrently, the microtubules come together to form bundles ( Figure 3 ). In the bundled state, cortical microtubules were preserved for up to 24 h while still in the plasmolytic medium. Interestingly, the cellulase KORRIGAN (KOR) showed similar microtubule arrays [ 38 ] but a role of microtubules in the transport of cellulose synthase (CESA) proteins from their origin in the Golgi towards the plasma membrane, e.g., by microtubule-associated CESA compartments (MASCs) or small CESA compartments (SmaCCs) is still under discussion (for a review, see [ 39 ]). Cortical microtubules labeled with GFP-MAP4 and GFP-TUA6 exhibit the same behavior under plasmolytic conditions but the background in GFP-TUA6 cells was higher ( Supplemental Figure S1 ). Therefore, only data for GFP-MAP4 are submitted hereafter. Microtubules or microtubule bundles were also present in Hechtian strands ( Figure 3 j). In deplasmolysis, microtubules remained organized in thick bundles, but they gradually separated from each other during the course of the protoplast swelling to its original size ( Supplemental Movie S1 ). In some samples, fluorescently labeled spots are observed ( Figure 3 h–j); these are located along microtubule bundles. It took up to 24 h until fine cortical microtubules were reinstated but even then, some thick tubular structures—probably corresponding to residual microtubule bundles—persisted (data not shown).

Green fluorescent protein (GFP)-tagged microtubules (GFP-MAP4) in Arabidopsis hypocotyl cells; ( a ) Interphase cells before plasmolysis; ( b ) Bundles and wavy microtubules in plasmolysed cells, treatment with 0.8 M mannitol for 30 min; bar: 10 µm.

Cortical microtubules (GFP-MAP4) in Arabidopsis hypocotyl cells during plasmolysis ( a – c ; f – j ) and deplasmolysis ( d , e ); plasmolysis/deplasmolysis times marked on the micrographs. ( a ) Bright field image showing cell walls, occasional stomata and chloroplasts (arrowheads); ( b ) Bundling of microtubules (arrows) at the onset of plasmolysis in 0.8 M mannitol; ( c ) After 40 min, cortical microtubules are showing waves and bundles; they remain aligned within the cortical cytoplasm; ( d ) Bundles of microtubules persist in deplasmolysis when the protoplasts become realigned at the cell wall; ( e ) Overlay of fluorescence and bright field image at the end of the plasmolytic cycle; chloroplasts appear as dark grey dots; ( f – j ) Higher magnification of plasmolysing hypocotyl cells; inlay in ( j ) shows microtubules in Hechtian strands (arrows). Bar: 10 µm.

2.3. Actin Microfilaments

Filamentous actin, as visualized by GFP-ABD in Arabidopsis hypocotyl cells, presented a fine network ( Figure 4 a). It consisted of radiating microfilaments and microfilament bundles which extended from the nuclear envelope towards the cortical parts of the cell ( Figure 4 b). Endoplasmic actin was present in transvacuolar cytoplasmic strands that traversed the vacuole thereby connecting the nuclear cytoplasm to the thin cortical cytoplasmic layer immediately below the plasma membrane. Since the subcortical cytoplasm was highly dynamic and organelles therein are constantly streaming in interphase cells, actin microfilaments needed to adjust accordingly. Therefore, it is easily understandable that actin microfilaments were able to reorganize rapidly in order to accommodate the shrinking protoplast during plasmolysis. The actin filaments radiating from the nuclear surface were still visible in plasmolysed cells ( Figure 4 b,c), while cortical actin lined the plasma membrane (arrowheads; Figure 4 c,d, inserts) and stretched out towards the cell wall in Hechtian strands ( Figure 4 f,g, arrows). The actin filaments did not become wavy as observed for microtubules. The plant cytoplasm contained a pool of globular actin in order to constantly assemble filaments; these in turn can rapidly disassemble if F-actin remodeling is necessary. Actin microfilament reorganization after hyperosmotic treatment was linked to Ca 2+ signaling and was associated with membrane mechanical integrity ([ 26 ] and references therein). However, it remains to be clarified if this Ca 2+ mobilization triggered actin filament formation directly by the increased polymerization of globular actin [ 40 ], or indirectly through a Ca 2+ signaling cascade [ 41 ].

Actin microfilaments (GFP-ABD) in Arabidopsis hypocotyl cells during a plasmolytic cycle; plasmolysis/deplasmolysis times marked on the micrographs. ( a , b ) Before plasmolysis, actin microfilaments were located in the cortical cytoplasm ( a ) and radiated from the nucleus ( b ); ( c ) In the plasmolysed state, actin microfilaments conserved this structure, but also lined the detached plasmalemma regions (arrows); ( d ) The protoplast rounded up at the onset of deplasmolysis, incorporating Hechtian strands (arrowheads) and ( e ) a “stabilizing layer” of bundled actin microfilaments followed the expanding protoplast; ( f , g ) High magnification of Hechtian strands (arrows) with actin microfilaments ( f ) and the corresponding bright field image ( g ), 30 min of plasmolysis in 0.8 M manntiol; arrowheads indicate the plasma membrane. Inserts ( c , d ): Single focal planes to show the cortical array. Bar: 10 µm.

At the onset of deplasmolysis, filamentous actin was clearly visible at the sites of big cytoplasmic aggregations: Around the nucleus and lining of the concave membrane-wall detachments. A weak fluorescent signal of GFP-ABD protein was observed along the plasma membrane of the expanding protoplast, even when there was only a very thin cortical layer. Hechtian strands became incorporated into the expanding protoplast. The cytoarchitecture at the start of deplasmolysis was preserved throughout the process ( Figure 4 d,e/ Supplemental Movie S2 ) until the protoplast was reconnected with the cell wall. Shortly after the plasmolytic cycle was completed, actin microfilaments in Arabidopsis hypocotyl cells were organized similarly to non-plasmolyzed control cells. Consequently, most filamentous actin was found to be present at sites of high cytoplasmic density as well as in the thin cortical layer below the plasma membrane. Organelle movement denoting cytoplasmic streaming was maintained throughout the whole plasmolytic cycle ( Supplemental Movie S3 ). In leaf cells of Chlorophytum comosum , the behavior of cortical actin filaments during plasmolysis has been described in terms of plasma membrane integrity and protoplast volume regulation [ 26 ]. In this study, actin microfilaments were shown to line the plasma membrane at areas of intense mechanical strain as in the case of concave plasmolysis. The authors report that many actin filament bundles formed a network, lining the areas of detached plasma membrane during concave plasmolysis. In addition, these fibers were compared to stress fibers found in animal cells. It is suggested that the formation of numerous cortical, subcortical and endoplasmic actin filaments was necessary to regulate shape and volume in plasmolysis (see also [ 42 , 43 ]). Furthermore, Komis and co-workers [ 26 ] describe the disappearance of thin cortical actin filaments during plasmolysis. This, however, could be attributed to the fact that cells were fixed and most of the actin in the cortical layer was lost. The GFP-tagged actin Arabidopsis line in the present study allowed for the use of living material. A faint fluorescence signal was observed along the protoplast, even at sites that contain hardly any cytoplasm.

2.4. A Broader View

Plasmolysis is not only used in laboratory experiments, it has been reported to occur naturally due to extracellular water withdrawal in freezing conditions [ 44 ]. A lot of current knowledge on the water balance of plants is based on extensive studies by Stadelmann [ 5 ] and his co-workers. In this study, although the focus was maintained on the cytoskeleton, keeping in consideration that it is part of a complex ER-cytoskeleton-plasma membrane-cell wall continuum, and plays an essential role in signaling and mechanosensing [ 21 , 45 , 46 ].

3. Experimental Section

Hypocotyl cells of Arabidopsis thaliana plants were used in this study. The plants were grown on ½ MS medium [ 47 ] under sterile culture conditions for five days. Apart from wild-type plants (Col 0), cytoskeletal elements were followed in green fluorescent protein (GFP)-tagged Arabidopsis lines. In order to visualize microtubules, Arabidopsis lines expressing a microtubule associated protein coupled to GFP (GFP-MAP4; [ 32 ]) were utilized; seeds were generously gifted by Professor Jozef Šamaj as well as an alpha tubulin encoding line (GFP-TUA6; [ 33 ]); kind gift from Dr. Sidney Shaw. Actin microfilaments were detected in Arabidopsis plants expressing a GFP-tagged actin binding domain of fimbrin 1 (GFP-ABD; [ 34 ]); another kind gift from Professor Jozef Šamaj.

Seedlings were secured between a microscope slide and a coverslip spaced by Parafilm stripes, and sealed using liquid petroleum jelly. The immobilization in the application of petroleum jelly prevented dislocation of the plantlets during liquid exchange and allowed for the observation of the same cells during a whole plasmolytic cycle. To induce plasmolysis, a 0.8 M mannitol solution was applied on one side of the coverslip and carefully removed from the opposite side using filter paper. Plasmolysis was completed after 30 to 40 min. Deplasmolysis was initiated by perfusing a 0.4 M mannitol solution for 15 min followed by the perfusion of distilled water.

Both plasmolysis and deplasmolysis, were observed under a confocal laser scanning microscope (Leica CTR SP5 linked to a DM6000CS stand) using a 63× water immersion lens. Z-stacks were taken at specific time frames (e.g., onset of plasmolysis, after 10 min, 30 min, 60 min, 12 h, and finally after 24 h). Time lapse video clips were also produced to follow the dynamic processes. For the videos shown as supplemental data , single images were taken at 2 min time intervals. The single image series were exported as .avi files by the Leica LASAF software at 12 fps. Editing was done with Adobe Premiere Pro version CS4 and CS6 (Adobe, San Jose, CA, USA).

4. Conclusions

Although plasmolysis is used in many cell biology experiments and student courses, the process itself and the incurred cytoarchitectural rearrangements remain to be fully understood. In this research paper, we describe cortical microtubule and actin microfilament organization during a plasmolytic cycle. Both structures are forced to adjust accordingly in the diminishing/expanding protoplast driven by vacuolar water efflux/influx. Further functional studies using stabilizing or disrupting agents on the cytoskeleton will allow for a more in-depth view on the role of cytoskeletal elements in plasmolysis.

Acknowledgments

This work was supported by a travel and accommodation grant (project INTERHANA, CZ.1.07/2.3.00/20.0165) to G.K. We gratefully acknowledge the gift of GFP-MAP4 and GFP-ABD lines by Jozef Šamaj (CR-Hana, Palacky University, Olomouc, Czech Republic) and of GFP-TUA6 by Sidney Shaw (Biology Department, Indiana University, Bloomington, IN, USA).

This article was supported by the Open Access Publishing Fund of the University of Vienna.

Supplementary Files

Supplementary file 1, author contributions.

I.L. initiated this contribution to the special issue on “Plant Vacuole” and wrote the manuscript with editorial assistance from S.S. and G.K. Plasmolysis experiments were performed by I.L., S.S. and G.K. B.S. edited the videos for supplemental data . All authors discussed and interpreted the results.

Conflicts of Interest

The authors declare no conflict of interest.

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Plasmolysis of red onion cells

Red onion cells were added with concentrated salt water. The cells started to lose water by osmosis. A picture was taken every 2 seconds to make a time lapse animation. Some cells do not contain a red pigment, because it was lost during the preparation of the cells.

Instructions can be found on page 8 of the January 2011 issue of the magazine: Microbehunter Magazine (January 2011)

2 thoughts on “Plasmolysis of red onion cells”

Simple trick, but looks really nice.Thanks for the instructions. Why did you make an animated gif and not a video? Greetings, Martin

A very interesting demonstration of plasmolysis reaction…

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Onion Cell Plasmolysis Lab

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Experiment on Plasmolysis (With Diagram)

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To demonstrate the phenomenon of plasmolysis.

Requirements:

Tradescantia leaf, water, sugar solution, slides, cover glass, microscope, blade.

1. From the lower surface of the leaf of Tradescantia, peel off small segments of epidermis by a blade.

2. Put few peelings on a slide, mount in a drop of water, put a cover glass and study under microscope.

3. Mount some of the peelings in the drops of sugar solutions of different concentrations. Study under the microscope.

Observations:

Peelings mounted in the water show clear cell structure. But the peelings placed in the sugar solution show the Concentra in their cell contents. More the concentration of sugar solution more is the contraction and shrinkage of cell contents. Peelings mounted in very high concentrated sugar solution, when observed under microscope, show complete shrinkage of their cell contents which become round or ball-like.

Shrinkage of the cell contents in the peelings mounted in conc. sugar solution is due to the fact that the osmotic pressure of the outer sugar solution is higher than that of the osmotic pressure of the cell sap. So the water from the cell sap diffuses into the external sugar solution through the semipermeable plasma membrane of the cell. Thus there is a shrinkage of the cell contents and this phenomenon is known as plasmolysis (Fig. 7).

If slightly plasmolysed (incipient plasmolysis.) cells are now kept in pure water, these will show the phenomenon of endosmosis and the cells will recover soon. This indicates the phenomenon of deplasmolysis.

Related Articles:

• Plasmolysis: Meaning and Importance | Plant Physiology
• Plasmolysis: Importance and Practical Utility of Plasmolysis (With Diagram)

Experiment , Biology , Physiology , Plasmolysis , Experiment on Plasmolysis

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Plasmolysis: Loss of Turgor and Beyond

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• Biology Article

Study Of Plasmolysis In Epidermal Peels

Table of contents, plasmolysis, materials required, observation, precautions, viva questions.

Plasmolysis is the process during, which cells lose water when placed in a hypertonic solution, that is greater in the concentration of solutes compared with the inside of a cell. During plasmolysis, the organelles inside the cell shrink away from the cell wall, which results in severe water loss and leads to the collapse of the cell wall and finally results in cell death.

Osmosis is primarily responsible for the occurrence of plasmolysis. There are three stages in plasmolysis. Based on certain criteria, plasmolysis can be classified into two different types:

• Concave plasmolysis – It is a reversible process.
• Convex plasmolysis- It is an irreversible process.

Also Refer:  Plasmolysis

Let us perform a simple experiment to study in detail the process of plasmolysis in the plant cell.

To study the phenomena of plasmolysis in epidermal peels of Rhoeo plant leaves in hypotonic and hypertonic solutions using salt solution.

• What is Plasmolysis?

It is a process of contraction or shrinkage of the protoplasm of a plant cell due to the loss of water from the cell. It takes place in extreme conditions and hence occurs rarely. Plasmolysis can be carried out in a laboratory by submerging a living cell in a concentrated sugar or salt solution for water loss from the cell.

• Why are the Rhoeo plant’s leaves are used in this experiment?

The Rhoeo plant’s leaves are used in this experiment because the Rhoeo leaf has coloured cell sap, which can be examined clearly under a compound microscope.

• How does water permeate through the cell membrane?

The cell membrane serves as a semipermeable membrane dividing the inner of all cells from that of its surroundings. This membrane permits movement of a few of the particles including, water molecules, and ions across the membrane while blocking others. There is the continuous movement of water molecules in and out of the cell across the cell membrane ad also it serves as an important attribute for enabling cells to absorb water.

Also Refer:   Cell Wall and Cell Membrane

• Glass slides.
• Watch glass.
• Rhoeo leaf.
• Coverslips.
• Compound microscope.
• Sodium chloride 5% solution.
• Sodium chloride 0.1% solution.
• Take two clean and dried glass slides and place them on a table.
• Select the fresh and cleaned Rhoeo leaves and place them on the watch glass.
• Fold the leaves in such a way that it tears from the lower side of the leaf. Or, with the help of a clean blade.
• Extract two small fragments of a fine and transparent layer with the help of forceps from the lower surface of the epidermis of the Rhoeo leaf.
• Now set up the epidermal peels on each of the glass slides.
• With the help of a dropper, add 1 to 2 drops of sodium chloride 0.1% solution to one of the prepared slides.
• With the help of another dropper, add 1-2 drops of sodium chloride 5% solution to the other prepared slide.
• Now carefully set a cover slip on the peel of both sides with the help of a needle. Make sure, no bubbles are present.
• Leave the prepared glass slide undisturbed for a few minutes.
• Now carefully place the slides under a compound microscope and observe the changes.

After a period of 30 minutes, we can notice that cells placed in the sodium chloride 0.1% solution seem to be turgid and the cells placed in the sodium chloride 5% solution seem to be shrunk with the loss of water and it exhibits the process of plasmolysis.

Plasmolysis is observed when the plant cells are immersed in the concentrated salt solution or sodium chloride 5%  solution. During this process, 4 to 5 per cent of water passes through the cell membrane into the encircling medium. This occurs as the concentration of water inside the cell is higher than the outside of the cell hence the protoplasm induces shrinkage and takes a spherical shape.

When the plant cells are immersed in a dilute salt solution or sodium chloride 0.1% solution, the water in the plant cells moves from the outside to the inside of the cell as the water concentration is higher outside the cell as compared to the inside of the cell which causes the turgidity of the cell.

Also Read:   Plant Water Relations

• The part of the Rhoeo leaf that needs to be extracted for the experiment is the epidermal peel from the lower surface
• Care needs to be taken to ensure that the peel is moist and not dry.

Q.1. What is Plasmolysis?

A.1. It is a process wherein the protoplasm of the plant cell turns rounded as a result of contraction when placed in a hypertonic solution due to exosmosis resulting in the decline in the tension of the cell wall.

Q.2. What is Incipient Plasmolysis?

A.2. It is the initial phase of plasmolysis wherein the protoplasm is just about to leave the cell wall.

Q.3. List any two significance of Plasmolysis? 

A.3. Listed below are the importance of plasmolysis :

• Helps to better understand and study the nature of a living cell
• It is used in the preservation of food materials such as jellies, and meat. Used in pickling as its salting is known to kill bacteria

Q.4. What is Osmotic pressure?

A.4. It is a pressure that checks the osmosis process. It needs to be applied to check the passage of the solvent as a result of osmosis.

Q.5. What is turgor pressure?

A.5 It is the pressure that presses the plasma membrane against the cell wall. It is a hydrostatic pressure that arises within the cell as a result of endosmosis on the cell wall.

Q.6. What is wall pressure?

A.6. The turgor pressure on the rigid walls of the cells causes exerting equal pressure in the opposing direction. That is termed wall pressure.

Q.7. When does the water potential become equal to that of its surroundings?

A.7. It happens when the turgor pressure becomes equal to the wall pressure thereby stopping the entry of water into the cells.

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