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Preparation of Onion Epidermal Cell Walls for Imaging by Atomic Force Microscopy (AFM)

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Nature Plants
Apr 2017



The growing plant cell wall is comprised of long, thin cellulose microfibrils embedded in a hydrated matrix of polysaccharides and glycoproteins. These components are typically constructed in layers (lamellae) on the inner surface of the cell wall, i.e., between the existing wall and the plasma membrane. The organization of these components is an important feature for plant cell growth and mechanics. To directly visualize the nano-scale structure of the newly-deposited surface of primary plant cell walls without dehydration or chemical extraction, a protocol of cell wall preparation for AFM imaging the most recently-synthesized cell wall surface in aqueous solutions was developed. Although the method was developed for onion scale epidermal peels, it can also be adapted to other organs, such as Arabidopsis hypocotyls, as well as ground samples of cell walls from the leaf petioles or hypocotyls of Arabidopsis and cucumber, maize coleoptiles and onion parenchyma. Potential artifacts of AFM imaging of plant cell walls are also discussed.

Keywords: Atomic force microscopy (原子力显微镜技术), Primary plant cell walls (初生植物细胞壁), Onion epidermis (洋葱表皮), Hypocotyls of Arabidopsis and cucumber (拟南芥和黄瓜下胚轴), Maize coleoptiles (玉米胚芽鞘)


The structure of primary plant cell walls plays a key role in determining cell wall biomechanical properties and regulating plant cell growth and morphogenesis (Cosgrove, 2005 and 2016). To visualize cell wall organization, common methods include transmission and scanning electron microscopy (TEM, SEM), light microscopy and AFM (McCann et al., 1990; Marga et al., 2005; Anderson et al., 2010; Ding et al., 2012; Abraham and Elbaum, 2013; Zhang et al., 2014; Zheng et al., 2017). In recent years AFM has enabled the imaging of soft biological samples in fluid, thus allowing studies of plant cell walls at nm-resolution in a close-to-native state, without dehydration, harsh chemical treatment, embedding or sectioning. Compared to EM techniques, which typically require dehydrated samples for high resolution imaging, AFM avoids dehydration artifacts and simplifies the sample preparation procedure while achieving high resolution imaging at the nanometer scale (Zhang et al., 2016). Combining AFM imaging with enzyme treatments, mechanical testing or use of cell wall mutants (Xiao et al., 2016; Zhang et al., 2017), we can test the hypothesized roles of specific cell wall components in cell growth, mechanics and cell wall structure.

For this protocol we focus on the onion scale epidermal cell wall, which has been the subject of numerous mechanical, spectroscopic and microscopic studies (Wilson et al., 2000; Kerstens et al., 2001; Hepworth and Bruce, 2004; Vanstreels et al., 2005; Loodts et al., 2006; Suslov et al., 2009). A key difference between our preparation method and that of previous authors is that our method splits open the epidermal cell layer, separating the outer epidermal cell wall from the remainder of the epidermal cell. After a brief wash to remove membranes and cellular debris, the newly-synthesized surface is ready for direct imaging by AFM. This is possible because the abaxial epidermis (on the convex or outer surface of the onion scale) adheres tightly to the underlying parenchyma tissues, so the peeling procedure splits open the epidermal cells. In previous studies with onion, whole-cell epidermal layers were peeled from the adaxial (inner or concave) surface of the onion scale, i.e., this cell layer adheres weakly to the underlying tissues and so separates intact at the interface (the middle lamella) with the underlying tissue. This difference in peeling procedure is of course important for surface-imaging methods such as AFM and SEM.

Materials and Reagents

  1. Razor blades
  2. Double sided tape (Permanent Double Sided Tape) (Scotch, catalog number: 3136 )
  3. Glass microscope slide (75 x 25 mm) (VWR, catalog number: 48300-025 )
  4. Charged microscope slide (75 x 25 mm) (VWR, catalog number: 48312-905 )
  5. 0.2 µm filter (Whatman GD/X 25 mm Sterile PVDF Syringe Filter) (GE Healthcare, catalog number: 6900-2502 )
  6. Fresh onions bulbs (Allium cepa, cv. Cometa), approximately 8 cm in diameter
  7. Nail polish (nitrocellulose dissolved in butyl and ethyl acetate)
  8. 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid sodium salt (HEPES) (Sigma-Aldrich, catalog number: H7006 )
  9. Tween-20 (Sigma-Aldrich, catalog number: P9416 )
  10. Sodium acetate, anhydrous (Sigma-Aldrich, catalog number: S2889 )
  11. Washing buffer (see Recipes)
  12. Imaging buffer (see Recipes)


  1. Forceps
  2. Bruker Dimension Icon atomic force microscope with ScanAsyst and PeakForce QNM (Quantitative Nanomechanical Property Mapping) operation package
  3. Scanasyst-fluid + AFM tips (Bruker, CA)
  4. Rocking platform (VWR, model: Model 200 )
  5. Slide warmer (50 °C) (Fisher Scientific, model: Model 77 , catalog number: 12-594)


  1. Nanoscope for AFM operation and Nanoscope Analysis for image analysis (Bruker)


  1. Purchase fresh onion bulbs (Allium cepa, cv. Cometa or similar white varieties, ~8 cm in diameter) from a local grocery store. The onions can be stored at 4 °C up to two weeks.
  2. Remove the dry, papery outer layers. The fleshy scales are numbered consecutively 1, 2, 3…, n with scale #1 being the oldest, outermost layer (Figure 1A).
  3. Excise scale #5 as a ~1 cm wide strip and press a 2 x 1.2 cm piece of double-sided tape to the convex, abaxial surface of the upper middle (near the tip) part of the onion scale (Figure 1B).
  4. Peel off the epidermis (with the tape) by hand, trim to ~1.5 x 1.2 cm (Figure 1C), and place the tape and the sample onto a clean glass slide, with the sample side facing up. Press the perimeter of the sample onto the slide.
  5. Seal the perimeter of the sample with a narrow line of nail polish and add a droplet (~20 μl) of washing buffer (see Recipes) to the center of the sample to keep it from drying (Figure 1D). The washing buffer should not submerge the surrounding still-wet nail polish. Steps 2 to 5 should be completed in ~30 sec.

    Figure 1. Procedure of onion epidermal wall preparation for AFM. A. Numbered onion scales with scale #1 being the oldest, outermost layer. Epidermal peels from scale #5 were usually used for imaging, although other scales are compatible with this procedure. B. A ~1 cm wide strip was excised and a double-sided tape was pressed against the abaxial side of the upper middle (near the tip) part of the onion scale. C. The epidermis was peeled off along with the tape. D. The epidermis and the tape were pressed onto the slide and a drop of washing buffer was added to the center of the sample.

  6. Allow the nail polish to cure for ~2 min or longer before adding more of the same washing buffer (~100 μl) and gently agitate on a rocking platform for at least 20 min. The slide is put in a Petri dish with lid closed to reduce evaporation. Before scanning, examine the peel under an optical microscope to verify that epidermal wall indeed separated from the rest of the cell, exposing the inner surface of the wall for direct contact with the AFM tip.
    Note: In our experience, peeling of the abaxial epidermis results in separation of the outer (periclinal) cell wall from the rest of the epidermal cell layer. However, this is not always uniform and must be verified by examination of the peel by optical microscopy after the detergent washing step. Cells with serrated edge or teeth-like features are usually torn open and good candidates for AFM imaging (Figure 2). The nucleus and other organelles should be absent. For AFM observations it is not essential that all of the cells split open in this way. Regions of the peel with intact cell walls can be ignored for the AFM imaging. The advantage of the onion scale abaxial epidermis is that it generally peels in this manner quite reproducibly and uniformly. Epidermal layers from other tissues sometimes split open in a similar fashion, others not. This must be determined by trial and error.

    Figure 2. Optical micrograph showing an AFM tip hovering above an epidermal cell. The serrated, teeth-like structures are good indications that the epidermal cells are split open.

  7. Calibrate the deflection sensitivity (~50-60 nm/V) and spring constant (0.2-0.7 N/m) of the AFM tip with a clean glass slide in fluid, following the manufacturer’s protocol. The tip radius (~1.8 nm), tip half angle (17.5°), and sample Poisson’s ratio (estimated as 0.3) need to be entered manually. The values given here are based on the tips that we routinely use (Scanasyst-fluid+ AFM tips) and may differ for other tips.
  8. Throughout the scanning processes, the samples are immersed in imaging buffer (20 mM HEPES buffer [pH 7.0] or 20 mM sodium acetate [pH 4.5], see Recipes).
  9. Start scanning at 2 x 2 μm size and 512 x 512 sampling rates (thus resolution of ~4 nm/pixel). For high resolution images, scan the sample at 0.5 x 0.5 μm or smaller. The scanning parameters are not identical for all materials, but typically fall in the following ranges, which are determined empirically: a peak force set point (600 pN-1.5 nN), scan rate (0.4-0.7 Hz), peak force frequency (1 kHz), tip velocity (0.9-1.5 µm/sec). The gain is auto controlled by the software but the set point and scan rate are typically controlled by the user for consistency throughout imaging (Figure 3).
  10. For standard cell wall imaging, scan at least five different cells from each sample and choose representative images for further analysis.

    Figure 3. A screenshot of Nanoscope interface during AFM operation. Four different channels are in display: Height, PeakForce Error, DMT modulus and Adhesion. Trace-retrace curves of each scan lines are displayed under each channel. The real-time force-distance curve is shown on the upper right corner.  

Data analysis

In Nanoscope Analysis software, the images are usually flattened to remove tilt or bow in each scan line and are exported in TIFF format. To export gray scale images, choose color table 0. For representative data see Zhang et al. (2014).


  1. All buffers must be of highest purity, using ddH2O and filtering through a 0.2 µm filter.
  2. Although much of our work is with onion scale #5, walls from other scales may also be used and studied in parallel with optical techniques (Kafle et al., 2014).
  3. While acquiring images, the real-time force curve should be monitored to optimize contact of tip to surface, hence generating a better-resolved image and more accurate mechanical mapping data. Also, the trace-retrace curves being displayed with each channel should be watched for indications of movement of the sample or poor tip-surface interaction. If the sample is mechanically unstable or if the tip does not make good contact with the surface, the resolution may be poor or imaging artifacts could be generated.
  4. To prepare walls from onion parenchyma, peel off the abaxial and adaxial side of epidermal layers, and grind the parenchyma to fine powder in liquid nitrogen. Wash the cell wall pellet with 20 mM HEPES buffer (pH 7.0) and 0.1% Tween-20 three times or until the supernatant is clear (centrifuge at 1,500 x g for 3 min). Resuspend the wall fragments in 20 mM HEPES buffer and add a droplet of the sample to a clean charged glass slide. Evaporate excess water on a slide warmer (50 °C) until the sample is still visibly moist but adheres to the slide (~15 min). Rinse the slide immediately with 20 mM HEPES buffer to remove loosely bound fragments before AFM scanning. We assume the middle lamella remains intact (Zamil et al., 2014) and so the only exposed wall surfaces are the inner faces of the cell walls. This assumption may not be true for other tissues.
  5. A similar grinding procedure can be used for Arabidopsis thaliana (Columbia) petioles (4-week-old; day/night, 16 h/8 h; temperature, 22 °C/16 °C), Arabidopsis dark grown hypocotyls (3 day at 22 °C), cucumber (Cucumis sativus) hypocotyls and maize (Zea mays) coleoptiles (dark grown for 4 days at 27 °C). Epidermal cell walls are identified by their characteristic appearance under the light microscope (elongated, rectangular shape, Figure 4). The cuticle side can be identified from the smoothness of the acquired image and the lack of fibrillar structures.

    Figure 4. A light micrograph showing the rectangular shaped epidermal fragment for AFM scanning

  6. Beware of possible artifacts induced by a dulled AFM tip or drifting due to temperature fluctuations or sample movement. The AFM tip can be contaminated or damaged during manufacturing process or during scanning, resulting in images with less sharp or ghosting features (Figure 5). Drifting is a common problem and should be examined in the beginning of the experiment by disabling ‘the slow axis scan’ (an option in scanning parameters) and repeatedly scanning one line. If drifting exists, the imaging features will appear tilted rather than vertically straight (Figure 6). Drifting can be limited by stabilizing samples in fluid for 1 or 2 h before collecting images. If the problem persists, scan a different cell or another sample.

    Figure 5. Common artifacts of AFM images due to dulled or broken tip. A. A representative AFM image of onion epidermis under normal conditions; B. Doubling/ghosting features of an AFM image due to a broken tip; C. Artefactual widening of microfibrils caused by a dulled tip.

    Figure 6. Disabling slow axis scan to check for drifting. A. Vertically straight features indicate there is no or very little drift. B. Tilted lines indicate drift.


  1. Washing buffer
    20 mM HEPES with 0.1% Tween-20, pH 7.0
  2. Imaging buffer
    20 mM HEPES (pH 7.0) or 20 mM sodium acetate (pH 4.5)


This protocol was adapted from previous work (Zhang et al., 2014; Zhang et al., 2016). We thank Ed Wagner and Liza Wilson for technical assistance and AFM maintenance and Dr. Sarah Kiemle for comments on the manuscript. This work was supported by the Center for Lignocellulose Structure and Formation, an Energy Frontier Research Center funded by the U.S. Department of Energy, Office of Science, Basic Energy Sciences (grant No. DE-SC0001090). The authors declare no conflict of interest.


  1. Abraham, Y. and Elbaum, R. (2013). Quantification of microfibril angle in secondary cell walls at subcellular resolution by means of polarized light microscopy. New Phytol 197(3): 1012-1019.
  2. Anderson, C. T., Carroll, A., Akhmetova, L. and Somerville, C. (2010). Real-time imaging of cellulose reorientation during cell wall expansion in Arabidopsis roots. Plant Physiol 152(2): 787-796.
  3. Cosgrove, D. J. (2005). Growth of the plant cell wall. Nat Rev Mol Cell Biol 6(11): 850-861.
  4. Cosgrove, D. J. (2016). Plant cell wall extensibility: connecting plant cell growth with cell wall structure, mechanics, and the action of wall-modifying enzymes. J Exp Bot 67(2): 463-476.
  5. Ding, S. Y., Liu, Y. S., Zeng, Y., Himmel, M. E., Baker, J. O. and Bayer, E. A. (2012). How does plant cell wall nanoscale architecture correlate with enzymatic digestibility? Science 338: 1055-1060.
  6. Hepworth, D. G and Bruce, D. M. (2004). Relationships between primary plant cell wall architecture and mechanical properties for onion bulb scale epidermal cells. J Texture Stud 35: 586-602.
  7. Kafle, K., Xi, X. N., Lee, C. M., Tittmann, B. R., Cosgrove, D. J., Park, Y. B. and Kim, S. H. (2014). Cellulose microfibril orientation in onion (Allium cepa L.) epidermis studied by atomic force microscopy (AFM) and vibrational sum frequency generation (SFG) spectroscopy. Cellulose 21: 1075-1086.
  8. Kerstens, S., Decraemer, W. F. and Verbelen, J. P. (2001). Cell walls at the plant surface behave mechanically like fiber-reinforced composite materials. Plant Physiol 127(2): 381-385.
  9. Loodts, J., Tijskens, E., Wei, C. F., Vanstreels, E., Nicolai, B. and Ramon, H. (2006). Micromechanics: Simulating the elastic behavior of onion epidermis tissue. J Texture Stud 37: 16-34.
  10. Marga, F., Grandbois, M., Cosgrove, D. J. and Baskin, T. I. (2005). Cell wall extension results in the coordinate separation of parallel microfibrils: evidence from scanning electron microscopy and atomic force microscopy. Plant J 43(2): 181-190.
  11. McCann, M. C., Wells, B. and Roberts, K. (1990). Direct visualization of cross-links in the primary plant cell wall. J Cell Sci 96: 323-334.
  12. Suslov, D., Verbelen, J. P. and Vissenberg, K. (2009). Onion epidermis as a new model to study the control of growth anisotropy in higher plants. J Exp Bot 60(14): 4175-4187.
  13. Vanstreels, E., Alamar, A. C., Verlinden, B. E., Enninghorst, A., Loodts, J. K. A., Tijskens, E., Ramon, H. and Nicolai, B. M. (2005). Micromechanical behaviour of onion epidermal tissue. Postharvest Biol Tec 37: 163-173.
  14. Wilson, R. H., Smith, A. C., Kacurakova, M., Saunders, P. K., Wellner, N. and Waldron, K. W. (2000). The mechanical properties and molecular dynamics of plant cell wall polysaccharides studied by Fourier-transform infrared spectroscopy. Plant Physiol 124(1): 397-405.
  15. Xiao, C., Zhang, T., Zheng, Y., Cosgrove, D. J. and Anderson, C. T. (2016). Xyloglucan deficiency disrupts microtubule stability and cellulose biosynthesis in Arabidopsis, altering cell growth and morphogenesis. Plant Physiol 170(1): 234-249.
  16. Zamil, M. S., Yi, H. and Puri, V. M. (2014). Mechanical characterization of outer epidermal middle lamella of onion under tensile loading. Am J Bot 101: 778-787.
  17. Zhang, T., Mahgsoudy-Louyeh, S., Tittmann, B. and Cosgrove, D. J. (2014). Visualization of the nanoscale pattern of recently-deposited cellulose microfibrils and matrix materials in never-dried primary walls of the onion epidermis. Cellulose 21: 853-862.
  18. Zhang, T., Vavylonis, D., Durachko, D. M. and Cosgrove, D. J. (2017). Nanoscale movements of cellulose microfibrils in primary cell walls. Nat Plants 3: 17056.
  19. Zhang, T., Zheng, Y. and Cosgrove, D. J. (2016). Spatial organization of cellulose microfibrils and matrix polysaccharides in primary plant cell walls as imaged by multichannel atomic force microscopy. Plant J 85(2): 179-192.
  20. Zheng, Y., Cosgrove, D. J. and Ning, G. (2017). High-resolution field emission scanning electron microscopy (FESEM) imaging of cellulose microfibril organization in plant primary cell walls. Microsc Microanal 23(5): 1048-1054.



【背景】原生植物细胞壁的结构在确定细胞壁生物力学性质和调节植物细胞生长和形态发生方面起着关键作用(Cosgrove,2005和2016)。为了显现细胞壁组织,常用方法包括透射和扫描电子显微镜(TEM,SEM),光学显微镜和AFM(McCann等人,1990; Marga等人, ,2005; Anderson等人,2010; Ding等人,2012; Abraham和Elbaum,2013; Zhang等人,,2014 ; Zheng等人,2017)。近年来,AFM已经使流体中的软生物样品成像,从而允许在接近天然状态下以毫微分辨率研究植物细胞壁,而不脱水,苛刻的化学处理,嵌入或切片。与通常需要脱水样品进行高分辨率成像的EM技术相比,AFM避免了脱水伪像,并简化了样品制备程序,同时在纳米级实现了高分辨率成像(Zhang等人,2016)。结合AFM成像与酶处理,机械测试或使用细胞壁突变体(Xiao等人,2016; Zhang等人,2017),我们可以测试假设特定细胞壁成分在细胞生长,力学和细胞壁结构中的作用。

对于该协议,我们关注洋葱鳞片表皮细胞壁,其已经成为许多机械,光谱学和显微镜研究的主题(Wilson等人,2000; Kerstens等人, ,2001; Hepworth和Bruce,2004; Vanstreels等人,2005; Loodts等人,2006; Suslov等人 ,2009)。我们的准备方法和以前的作者之间的一个主要区别是我们的方法分裂开表皮细胞层,从表皮细胞的其余部分分离外表皮细胞壁。经过短暂的清洗以去除膜和细胞碎片后,新合成的表面准备好通过AFM直接成像。这是可能的,因为离轴表皮(在洋葱鳞片的凸面或外表面上)紧密地粘附在下面的薄壁组织上,所以剥离程序分裂开表皮细胞。在以前对洋葱的研究中,全细胞表皮层从洋葱鳞片的正面(内部或凹陷)表面剥落,这种细胞层粘附到下面的组织上,因此完全分离界面(中间薄片)与下面的组织。这种剥离程序的差异对于诸如AFM和SEM的表面成像方法当然是重要的。

关键字:原子力显微镜技术, 初生植物细胞壁, 洋葱表皮, 拟南芥和黄瓜下胚轴, 玉米胚芽鞘


  1. 剃刀片
  2. 双面胶带(永久双面胶带)(Scotch,目录号:3136)
  3. 玻璃显微镜幻灯片(75×25毫米)(VWR,目录号:48300-025)
  4. 充电显微镜幻灯片(75×25毫米)(VWR,目录号:48312-905)
  5. 0.2微米过滤器(Whatman GD / X 25毫米无菌PVDF注射器过滤器)(GE Healthcare,目录号:6900-2502)
  6. 新鲜洋葱鳞茎(洋葱cepa ,cv。Cometa),直径约8厘米
  7. 指甲油(硝酸纤维素溶于丁基和乙酸乙酯)
  8. 4-(2-羟乙基)哌嗪-1-乙磺酸钠盐(HEPES)(Sigma-Aldrich,目录号:H7006)
  9. 吐温-20(Sigma-Aldrich,目录号:P9416)
  10. 乙酸钠,无水(Sigma-Aldrich,目录号:S2889)
  11. 洗涤缓冲液(见食谱)
  12. 成像缓冲区(见食谱)


  1. 镊子
  2. 布鲁克尺寸图标原子力显微镜与ScanAsyst和PeakForce QNM(定量Nanomechanical属性映射)操作包
  3. Scanasyst-fluid +原子力显微镜(布鲁克,加州)
  4. 摇摆平台(VWR,型号:200型)
  5. 幻灯片加热器(50°C)(Fisher Scientific,型号:77型,目录号:12-594)


  1. 用于AFM操作的纳米镜和用于图像分析的纳米分析(Bruker)


  1. 从当地的杂货店购买新鲜的洋葱鳞茎(洋葱cepa ,cv。Cometa或类似的白色品种,直径〜8厘米)。洋葱可以在4°C储存两周。
  2. 去除干燥的纸质外层。肉质鳞片连续编号为1,2,3 ...,n,鳞片#1是最老的最外层(图1A)。
  3. 将5号刻度作为〜1厘米宽的条带,并将2×1.2厘米的双面胶带压在洋葱刻度的上中部(靠近尖端)的凸面,背面上(图1B)。
  4. 用手剥去表皮(用胶带),修剪至〜1.5×1.2厘米(图1C),并将胶带和样品放在干净的载玻片上,样品面朝上。按下样品的周长到幻灯片上。
  5. 用一条细小的指甲油密封样品的周边,在样品中心加一滴水(〜20μl)清洗缓冲液(见配方),以防止其干燥(图1D)。洗涤缓冲液不应淹没周围静止的指甲油。步骤2至5应在约30秒内完成。

    图1.原子力显微镜下洋葱表皮的制备步骤A.刻度#1的洋葱鳞片是最老的最外层。通常使用5号刻度的表皮来进行成像,但是其他的刻度与这个程序是一致的。 B.切下1厘米宽的条状物,并将双面胶带压在洋葱鳞上部中部(尖端附近)的远轴侧。 C.表皮与胶带一起剥离。 D.将表皮和胶带压在载玻片上,并将一滴洗涤缓冲液加到样品的中心。

  6. 在加入更多相同的洗涤缓冲液(〜100μl)之前,让指甲油固化约2分钟或更长时间,并在摇摆平台上轻轻摇动至少20分钟。将玻片置于盖子关闭的培养皿中以减少蒸发。在扫描之前,检查光学显微镜下的剥离,以确认表皮细胞壁确实与细胞的其余部分分开,暴露出壁的内表面直接与AFM尖端接触。
    注意:根据我们的经验,离轴表皮的剥离导致外(周边)细胞壁与表皮细胞层的其余部分分离。然而,这并不总是一致的,并且必须通过在洗涤剂清洗步骤之后通过光学显微镜检查剥离来验证。具有锯齿状边缘或类似牙齿特征的细胞通常是开放的,并且是AFM成像的良好候选者(图2)。应该没有核和其他细胞器。对于原子力显微镜的观察,并不是所有的细胞都以这种方式分裂是不必要的。 AFM成像可以忽略具有完整细胞壁的果皮区域。洋葱鳞片表皮的优点是它通常以这种方式非常可重复和均匀地剥离。来自其他组织的表皮层有时会以相似的方式分裂,其他组织则不会。这必须通过反复试验来确定。

    图2.光学显微照片显示AFM尖端盘旋在表皮细胞上方。 锯齿状的牙齿状结构是表皮细胞分裂开的好迹象。

  7. 根据制造商的协议,用干净的载玻片将AFM尖端的偏转灵敏度(〜50-60 nm / V)和弹簧常数(0.2-0.7 N / m)校准。需要手动输入尖端半径(〜1.8nm),尖端半角(17.5°)和样品泊松比(估计为0.3)。这里给出的数值是基于我们经常使用的提示(Scanasyst-fluid + AFM提示),对于其他提示可能会有所不同。
  8. 在整个扫描过程中,将样品浸入成像缓冲液(20mM HEPES缓冲液[pH 7.0]或20mM乙酸钠[pH 4.5]中,参见食谱)。
  9. 以2 x 2μm大小和512 x 512采样率开始扫描(因此分辨率约为4 nm /像素)。对于高分辨率的图像,扫描0.5 x 0.5μm或更小的样品。所有材料的扫描参数并不相同,但通常落入以下经验确定的范围:峰值力设定点(600pN-1.5nN),扫描速率(0.4-0.7Hz),峰值力频率(1 kHz),尖端速度(0.9-1.5μm/ sec)。增益由软件自动控制,但设定值和扫描速率通常由用户控制,以保证整个成像的一致性(图3)。
  10. 对于标准细胞壁成像,扫描每个样品至少5个不同的细胞,并选择代表性的图像进行进一步分析。

    图3. AFM操作期间Nanoscope界面的屏幕截图。显示四个不同的通道:高度,峰值误差,DMT模量和粘附力。每条扫描线的曲线回扫曲线显示在每个通道下。实时力 - 距离曲线显示在右上角。  


在Nanoscope Analysis软件中,图像通常是平坦的,以消除每个扫描线上的倾斜或弯曲,并以TIFF格式输出。要输出灰度图像,请选择颜色表0.有代表性的数据请参见Zhang et al。 (2014)。


  1. 所有的缓冲液必须纯度最高,使用ddH 2 O和0.2μm过滤器过滤。
  2. 虽然我们的大部分工作是洋葱5号标尺,但是也可以使用其他尺度的墙壁,并与光学技术同时进行研究(Kafle et。,2014)。
  3. 在获取图像时,应监测实时力曲线以优化针尖与表面的接触,从而生成更好的分辨图像和更精确的机械制图数据。此外,应该观察每个通道显示的跟踪 - 回扫曲线,以了解样本移动的指示或较差的端面相互作用。如果样品机械不稳定或尖端与表面没有良好接触,则分辨率可能较差或可能会产生成像伪影。
  4. 为了从洋葱薄壁组织中制备壁,剥离表皮层的背面和近轴面,并在液氮中研磨薄壁细粉。用20mM HEPES缓冲液(pH7.0)和0.1%Tween-20洗涤细胞壁沉淀三次,或直到上清液澄清(在1500gxg离心3分钟)。重悬在20毫米HEPES缓冲区的墙壁碎片,并添加一个样品液滴到一个干净的带电玻璃幻灯片。在一台载玻片加热器(50°C)上蒸发多余的水分,直到样品仍然明显湿润,但粘附在载玻片上(约15分钟)。在AFM扫描之前,立即用20mM HEPES缓冲液冲洗载玻片以除去松散结合的片段。我们假设中间的薄片保持完好(Zamil et al。,2014),所以唯一暴露的壁表面是细胞壁的内表面。这个假设对于其他组织来说可能不是这样。
  5. (哥伦比亚)叶柄(4周龄,白天/夜间,16小时/ 8小时,温度,22℃/ 16℃),可以使用类似的研磨程序, (22℃下3天),黄瓜(Cucumis sativus )下胚轴和玉米( Zea mays )胚芽鞘(暗生长的在27℃4天)。在光学显微镜下,表皮细胞壁通过其特征外观来识别(细长的矩形形状,图4)。


  6. 注意可能由于温度波动或样品移动而导致钝角AFM尖端或漂移引起的伪影。在制造过程中或在扫描过程中,AFM针尖可能被污染或损坏,从而导致图像的锐利度较低或出现重影(图5)。漂移是一个常见的问题,应该在实验开始时通过禁用“慢轴扫描”(扫描参数选项)和反复扫描一行来检查。如果存在漂移,则成像特征将呈现倾斜而不是垂直笔直(图6)。在收集图像之前,漂浮可以通过在流体中稳定样品1或2小时来进行限制。如果问题仍然存在,请扫描其他单元格或其他样本。

    图5.由于尖端钝化或断裂导致的AFM图像的常见伪影。 :一种。洋葱表皮在正常条件下的代表性原子力显微镜图像; B.由于尖端断裂而导致AFM图像的倍增/重影特征; C.由一个钝的尖端引起的微纤维的人为扩大。

    图6.禁用慢轴扫描以检查漂移。 :一种。垂直笔直的特征表明没有或很少漂移。 B.倾斜的线条表示漂移。


  1. 清洗缓冲液
    具有0.1%Tween-20,pH 7.0的20mM HEPES
  2. 成像缓冲区
    20 mM HEPES(pH 7.0)或20 mM乙酸钠(pH 4.5)


该协议是从以前的工作改编的(Zhang等人,2014; Zhang等人,2016年)。我们感谢Ed Wagner和Liza Wilson提供技术援助和AFM维护以及Sarah Kiemle博士对手稿的评论。这项工作得到了由美国能源部科学基础能源科学局资助的能源前沿研究中心木质纤维素结构和形成中心(批准号:DE-SC0001090)的支持。作者宣称没有利益冲突。


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引用:Zhang, T. and Cosgrove, D. J. (2017). Preparation of Onion Epidermal Cell Walls for Imaging by Atomic Force Microscopy (AFM). Bio-protocol 7(24): e2647. DOI: 10.21769/BioProtoc.2647.