Quantifying the Permeability of the Apoplastic Water Barrier in Cosmos Petals

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The capacity of plants to minimize uncontrolled water loss is essential for survival in adverse and changing climatic conditions. In order to assess and compare the effectiveness of apoplastic barriers to water, the permeability of the barrier must first be quantified. Studies have accomplished this directly by quantifying tritium flux or indirectly by measuring the influx/efflux of water surrogates such as dyes, chlorophyll, and herbicides. Other studies have relied on comparative methods such as survival rates after drought. These methods rely on radioactive material, correlations, or qualitative comparisons. However, a quantitative method is necessary that directly measures water efflux and that allows easy comparisons within and between experiments, plant parts, plant species, and especially research laboratories. Here we outline in detail a gravimetric protocol first described by Schönherr and Lendzian (1981) that can be set up in less than half a day and completed in one to ten days depending on the plant barrier. This approach has been used in numerous studies on leaf and fruit cuticles and recently also on petals from cosmos (Cosmos bipinnatus; Buschhaus et al., 2015).

Materials and Reagents

  1. Leaf punch (12 millimeter diameter)
  2. Syringe (10 ml)
  3. Syringe (1 ml) with needle
  4. Pipette tip (100 µl)
  5. Pencil (unused)
  6. Sand paper (80 grit) (other grits may work but the author has not tested them)
  7. Tape (3M, Scotch®)
  8. Silica desiccant (Sorbsil Chameleon, BDH Prolabo)
  9. Plant material with minimum dimensions of 13 x 13 millimeters (e.g., ray petals 3-days post-anthesis from Cosmos bipinnatus cv. sensation pinkie; Seeds were obtained from Stokes Seeds)
  10. Silicon grease (Dow Corning Corporation, catalog number: Z273554 )
  11. De-ionized water
  12. Fertilizer (e.g., MiracleGro)


  1. Cylinders (machined in-house with an 11 mm diameter opening and an internal capacity of at least 1 ml; Figure 1. Further details are provided in the ‘Notes’ section)
  2. Metal meshing (10 cm x 20 cm x 0.2 cm; 0.7 cm hole diameter; 0.5 holes cm-2)
  3. Plastic box with tight fitting lid (10 cm x 20 cm)
  4. Incubator (Thermo Fisher Scientific, model: Isotemp 210 )
  5. Balance (Sartorius AG, model: CP1245 )
  6. Computer


  1. Spreadsheet software (e.g., Microsoft Excel)
  2. Data logger program to transfer balance data directly to computer (optional)


  1. Set an incubator to 25 °C. Allow sufficient time (e.g., overnight in our case) for the temperature to stabilize before plant material is harvested.
  2. Warm n+1 ml of water to 25 °C, where n equals the number of samples to be analyzed. We place a bottle of water in the incubator when we first turn it on.
  3. Fill the plastic container to a depth of approximately 1 cm with dry desiccant (Figure 1). Place the metal meshing on top of the desiccant to provide a solid base for the cylinders. Put the lid on the plastic container and then pre-warm to 25 °C in the incubator. We warm the container overnight just like the water in step 2, but one hour is likely sufficient. If the incubator does not occlude light, ensure that the selected plastic container does not transmit light.

    Figure 1. Cross-sectional diagram of the experimental set-up. Desiccant (10) is layered in to the bottom of a plastic container (11) that has a tight-fitting lid (1). A firm mesh surface (9) provides support for the cylinder (4). A petal disc (8) is sealed with silicon grease (7) in the 1.1 cm opening of the cylinder. Tape (6) provides an additional seal and holds the two parts of the cylinder together. The central opening in the chamber (5) is filled with water through a tiny hole (3) before the hole is sealed with tape (2).

  4. Assemble a ‘sealant applicator’ (Figure 2A). Screw a 100 µl pipette tip into the end of a plastic 10 ml syringe. Cut the end of the pipette tip to the desired diameter, which in our case is 1.5-2 mm. Remove the plunger and fill the syringe with a sealant such as high temperature vacuum grease. Re-insert the plunger. Slowly pressing the plunger allows for a thin bead of sealant to be evenly and continuously applied to the chambers. Other sealants such as Vaseline are likely sufficient; the authors however have not experimented with this.

    Figure 2. Tools used in this protocol. A sealant applicator was constructed by screwing a 100 µl pipette tip into the end of a disposable syringe (A). The syringe is then filled with sealant. Since the pipette tip is tapered, the diameter of the applied sealant bead can be modified by trimming the pipette tip. A surface abrader was constructed by gluing a small disc of 80 grit sand paper to the bottom of a pencil (B). A 12 mm or 13 mm leaf punch was used to cut a disc from the petal (C).

  5. Prepare the cylinders by applying a bead of sealant (silicon grease) using the ‘sealant applicator’ in a continuous, single ring on the inner surface of the cylinder lid and in a continuous, single ring on the cylinder bottom where it will contact the lid (Figure 4A). The sealant will be spread out later (steps 10-11) in the process of joining together the two surfaces to be sealed. Pre-warm the cylinders to 25 °C by putting them in the incubator.
  6. Construct a ‘surface abrader’ (Figure 2B). Glue a circle of sand paper on to the eraser end of a new pencil. Try using a hole punch to obtain a suitably sized circle of sand paper.
  7. Obtain plant material - cosmos (Cosmos bipinnatus cv. Sensation pinkie) ray petals from flowers 3-7 days post anthesis - with dimensions greater than 13 mm by 13 mm (Figure 3A). Controlled conditions of plant growth aid in reproducibility. Astomatous leaves of other plants may also be used, provided that they have an area greater than 13 mm by 13 mm. If the adaxial and abaxial surfaces are indistinguishable, mark the opposing surface with a permanent marker (Figure 3B); do not mark the surface of interest.

    Figure 3. Preparation of petal disc for water permeance analysis. A ray petal is harvested from a flower three days post anthesis (A). The side opposite from the surface-of-interest is marked in situations where the adaxial surface is indistinguishable from abaxial surface (B). The surface abrader is gently twirled on this opposite surface (C) to scratch through the surface such that water can more easily penetrate from this side (D). A cork borer is used to cut the petal (E) into a disc, ready for sealing into the cylinder (F).

  8. Gently abrade the opposing, marked surface (not the surface of interest) with a gentle rotation of the sandpaper-end of the ‘surface abrader’ (Figure 3C-D). The purpose of this step is to facilitate water ingress, thereby ensuring a maximal internal water concentration. Failure to disturb the cuticular barrier on this surface will invalidate the water loss measurements. A razor blade may be used to cut slits instead of abrasion with the sandpaper. However, this requires steady hands and/or thick plant material to prevent slicing through the cuticle of interest.
  9. Using the leaf punch (cork borer; Figure 2C), stamp a disc of plant tissue measuring 12 mm in diameter (Figure 3E-F).
  10. Place the plant disc into the lid such that the surface-of-interest faces the out towards the desiccant (Figure 4B-C). Use the ‘surface abrader’ to gently tap around the disc circumference such that the disc makes continuous contact with the sealant bead.
  11. Gently slide the chamber bottom onto the lid. Rotate the bottom a quarter turn to ensure the sealant is evenly spread between the cylinder lid and bottom.
  12. Wrap a strip of tape around the cylinder such that it overlaps both the lid and the bottom. This provides a secondary seal as well as prevents the lid and bottom from separating in subsequent handling (Figure 4D-E).
  13. Fill the cylinder with 1 ml of the pre-warmed water through the small opening in the bottom of the cylinder (Figure 1 Item 3). We used a syringe to facilitate this transfer. However, do not insert the syringe needle so far into the hole that it punctures the plant disc. Also, gently expel the water so as not to disturb the sealed petal disc. Seal the water-filling hole with a piece of clear tape.
  14. Label the cylinder. We number (or letter) our cylinders and record this number with a description of the samples, including plant species, plant part, surface, age of specimen, treatment, and date.
  15. Place the cylinder in the plastic container and return to the incubator.
  16. Repeat steps 8-15 for each sample to be analyzed.
  17. Leave the cylinders in the desiccant container in the 25 °C incubator overnight to stabilize.
  18. Weigh the cylinders, recording the time of measurement and mass, and return them to the desiccant container and incubator. Although optional, connecting the balance to a computer via a data logging program to record the time and mass in a Microsoft Excel-compatible file format simplifies data handling. It is assumed that prior to weighing the cylinders the balance has been calibrated according to the manufacturer’s instructions. We recommend also weighing a standard of similar mass to the samples. This allows the accuracy of the balance to be confirmed when the measurements are made over several days (see following step).
  19. Repeat step 18 at set time intervals. The elapsed time must allow sufficient water loss such that the new mass decreases from the previous mass by more than the error of the balance. For fairly permeable plant surfaces, e.g., cosmos petals (Buschhaus et al., 2015), a measurement every 2 h is adequate since, for our balance, d = 0.1 mg. However, samples are typically weighed once per day to decrease the percent error. More water-resistant plant surfaces may require even longer intervals than this. The time interval may be increased or decreased to account for the precision of your balance.
    While each cylinder is removed for weighing, inspect the colour of the desiccant underneath. If the desiccant immediately under the cylinder has changed to the hydrated colour, the plant disc has likely ruptured. Gently turn the cylinder over and inspect the plant disc for macroscopic tears and discard torn samples.
  20. Plot the cumulative change in mass per cumulative change in time. The linear regression line typically accounts for greater than 97% of the observed variation. A quick examination of the plot when r2 values are less than 0.97 usually reveals a distinct bend in the line at a point where either the plant disc ruptured or the chamber ran out of water. Representative data can be found in Figure 2 of Buschhaus et al. (2015).
  21. Calculate the permeance P (m s-1) according to the equation P = F A-1 C-1. F is the flow (mg s-1) and equals the slope of the linear regression line from the previous step. A is the exposed area of the leaf disc. Ensure that this is the true surface area and not just the apparent surface area. This is especially important for the adaxial surface of petals where the epidermal cells frequently are not flat. C is the change in water vapour concentration (23.07 mg m-3 at 25 °C). Convention dictates that the geometric mean is reported as the data are typically skewed towards higher permeances (Baur, 2008). For a further explanation of the equation, see Schreiber and Schönherr (2009).

    Figure 4. Cylinder for the gravimetric measurement of water permeance. Silicon grease (white bands) is applied in a ring on the lid and on the bottom of the cylinder (A). The petal disc is placed between the openings in the lid and bottom (B) and the lid is pressed firmly onto the bottom (C). The lid and bottom are secured together with a strip of tape (hatched white; D) to complete the cylinder assembly (E).


  1. The dimensions and composition of the cylinder are adaptable. For ease of subsequent calculations, the opening into the center of the cylinder should have a constant diameter. We find that a diameter of 11 mm was satisfactory for Cosmos petals and also allowed the cylinders to be used for the analysis of leaves from other species. The internal volume should hold at least 1 ml of water as this permits measurable water loss (up to a 1 g change-in-mass for a balance that reads to 0.0001 g) before running out. A 2 mm diameter hole was drilled in the bottom to allow the cylinder to be filled with water after the petal was sealed across the opening. In our case, the cylinders were machined out of stainless steel. However, any material not permeable to water should be adequate. External dimensions were approximately 2.5 cm by 2 cm (diameter by height).
  2. In order to test your sealing technique, seal a penny across the opening of the cylinder instead of the plant disc. With this set-up, no change in mass should be observed.
  3. Although not an issue for petals (or astomatous leaves), many leaves have stomates that provide an alternative route for water movement through the barrier. The cylinders may be incubated under conditions that promote stomate closure but care must still be exercised in interpreting the results.
  4. Several variations can be explored with this experimental set-up. After quantifying the barrier in a plant disc, the epicuticular wax layer can be removed and the disc re-analyzed. Following this, all of the cuticular wax can be removed and the disc re-analyzed. In this way the relative contributions of the cuticular wax layers can be determined.
  5. Easiest plant growth occurs with direct seeding of Cosmos on to soil (e.g., Sunshine mix 4; 1 plant per 15 cm diameter pot), weekly watering with fertilizer, and growth conditions of constant 20 °C temperature under 12 h day (approx. 150 µmol m-2 s-1 photosynthetically active radiation)/12 night conditions. If space is limited, seeds can first be sprouted prior to transplanting to soil as documented in Buschhaus et al. (2015) to ensure that every pot has a successfully growing plant.


The authors thank Dr. L. Schreiber (University of Bonn, Germany) for suggesting the construction of the surface abrader. The methods used in this publication were first described in Schönherr and Lendzian (1981) and first adapted to petals by Buschhaus et al. (2015). Generous support to R. Jetter has been provided by the Natural Sciences and Engineering Research Council (Canada), the Canada Research Chairs Program, and the Canada Foundation for Innovation.


  1. Baur, P. (2008). Lognormal distribution of water permeability and organic solute mobility in plant cuticles. Plant Cell Environ 20(2): 167-177.
  2. Buschhaus, C., Hager, D. and Jetter, R. (2015). Wax layers on Cosmos bipinnatus petals contribute unequally to total petal water resistance. Plant Physiol 167(1): 80-88.
  3. Schreiber, L. and Schönherr, J. (2009). Water and solute permeability of plant cuticles. Springer-Verlag Berlin Heidelberg.
  4. Schönherr, J. and Lendzian, K. (1981). A simple and inexpensive method of measuring water permeability of isolated plant cuticular membranes. Z Pflanzenphysiol 102(4): 321-327.


植物使不受控制的水损失最小化的能力对于在不利和变化的气候条件下的存活是必要的。为了评估和比较非水性阻隔层对水的有效性,必须首先量化阻隔层的渗透性。研究直接通过量化氚通量或通过测量水替代物如染料,叶绿素和除草剂的流入/流出间接实现了这一点。其他研究依赖于比较方法,如干旱后的存活率。这些方法依赖于放射性材料,相关性或定性比较。然而,定量方法是必要的,直接测量水流出,并允许在实验,植物部分,植物物种,特别是研究实验室内和之间的简单比较。这里我们详细介绍了Schönherr和Lendzian(1981)首次描述的重量分析方法,它可以在不到半天内完成,并且根据植物屏障在一到十天内完成。这种方法已经用于许多关于叶和果皮表皮的研究中,并且最近也用于来自宇宙的花瓣( Cosmos bipinnatus ; Buschhaus ,2015)。


  1. 叶冲(直径12毫米)
  2. 注射器(10ml)
  3. 注射器(1 ml)用针
  4. 移液器吸头(100μl)
  5. 铅笔(未使用)
  6. 砂纸(80粒度)(其他砂粒可能工作,但作者没有测试它们)
  7. 胶带(3M,Scotch )
  8. 二氧化硅干燥剂(Sorbsil Chameleon,BDH Prolabo)
  9. 植物材料,具有13×13毫米的最小尺寸(例如射线花瓣从开花后3天天花来自Cosmos bipinnatus cv。感觉小指;种子获自Stokes Seeds) br />
  10. 硅油(Dow Corning Corporation,目录号:Z273554)
  11. 去离子水
  12. 肥料(例如 MiracleGro)


  1. 气瓶(内部加工,直径为11 mm,内部容积至少为1 ml;图1.更多详细信息,请参见"注意事项"部分)
  2. 金属网格(10cm×20cm×0.2cm; 0.7cm孔径; 0.5孔cm <-2μm)
  3. 塑料盒带紧密盖(10厘米x 20厘米)
  4. 孵育器(Thermo Fisher Scientific,型号:Isotemp 210)
  5. 平衡(Sartorius AG,??型号:CP1245)
  6. 电脑


  1. 电子表格软件(例如 Microsoft Excel)
  2. 数据记录器程序将平衡数据直接传送到计算机(可选)


  1. 将孵育器设置为25°C。在收获植物材料之前,让温度稳定,以留出足够的时间(例如在我们的情况下过夜)。
  2. 温热n + 1ml水至25℃,其中n等于待分析的样品数。当我们第一次打开它时,我们在培养箱中放一瓶水
  3. 用干燥的干燥剂填充塑料容器至约1cm的深度(图1)。将金属网放在干燥剂的顶部,为圆柱体提供坚固的基底。将盖子放在塑料容器上,然后在孵化器中预热至25°C。我们像在步骤2中的水一样温暖容器,但是一小时可能就足够了。如果孵育器没有遮光,请确保所选的塑料容器不透光。


  4. 组装"密封胶涂布器"(图2A)。将100μl移液器吸头拧入塑料10ml注射器的末端。将移液管末端切成所需的直径,在本例中为1.5-2 mm。取下柱塞,用高温真空润滑脂等密封剂填充注射器。重新插入柱塞。缓慢地按压柱塞允许密封剂的小珠均匀且连续地施加到腔室。其他密封剂如凡士林可能就足够了;但作者没有尝试这一点。

    图2.本协议中使用的工具。通过旋拧100μl移液管尖端 进入一次性注射器(A)的端部。然后填充注射器 与密封剂。由于移液管尖端是锥形的,直径 应用的密封胶珠可以通过修剪移液器吸头修改。一个 通过胶合80砂粒的小圆盘来构造表面磨损器 纸到铅笔底部(B)。 12mm或13mm的叶冲头 用于从花瓣(C)切割光盘。

  5. 通过在气缸盖内表面上的连续单环中使用"密封胶涂布器"施加密封剂珠(硅润滑脂),并在气缸底部上的连续的单个环中施加气缸以制备气缸,在那里它将接触盖(图4A)。在将要密封的两个表面连接在一起的过程中,密封剂将在以后铺开(步骤10-11)。将气瓶放入培养箱中,将气瓶预热至25°C
  6. 构造"表面磨损"(图2B)。将一圈沙纸粘贴到新铅笔的橡皮擦端。尝试使用打孔机获得合适尺寸的砂纸。
  7. 在开花后3-7天从花中获得植物材料 - 宇宙( cv。感觉小指)光线瓣 - 尺寸大于13mm×13mm(图3A)。植物生长的控制条件有助于再现性。也可以使用其它植物的气味叶,只要它们的面积大于13mm×13mm。如果近轴和远轴表面不可区分,用永久标记标记相对的表面(图3B);不要标记感兴趣的表面

    图3.用于水渗透性分析的花瓣瓣的制备。开花后三天从花中收获射线花瓣(A)。 在情况中标记与感兴趣的表面相对的一侧 其中近轴表面与背轴表面(B)不可区分。 ?表面研磨器在该相对表面(C)上轻轻地旋转 划伤通过表面使得水可以更容易地渗透 从该侧(D)。软木钻孔器用于将花瓣(E)切成 盘,准备用于密封到气缸(F)中
  8. 用"表面研磨器"的砂纸端轻轻旋转轻轻磨擦相对的,标记的表面(不是感兴趣的表面)(图3C-D)。该步骤的目的是促进水进入,从而确保最大内部水浓度。不能打扰表面上的表皮屏障将使水损失测量失效。剃刀刀片可以用于切割狭缝而不是用砂纸磨损。然而,这需要稳定的手和/或厚植物材料以防止通过感兴趣的角质层切片
  9. 使用叶冲头(软木钻孔器;图2C),压印直径为12mm的植物组织的盘(图3E-F)。
  10. 将植物圆片放入盖子,使感兴趣的表面朝向干燥剂的外面(图4B-C)。使用"表面磨损工具"轻轻敲击圆盘周围,使圆盘与密封胶珠连续接触。
  11. 轻轻地将瓶底滑到盖子上。将底部旋转四分之一圈,以确保密封胶均匀地分布在缸盖和底部之间
  12. 将一条胶带缠绕在圆筒上,使其与盖子和底部重叠。这提供了二次密封,并且防止在随后的处理中盖子和底部分离(图4D-E)
  13. 通过汽缸底部的小开口向汽缸中加入1毫升预热的水(图1项目3)。我们使用注射器来促进这种转移。但是,请勿将注射器针头插入到足以刺穿植物圆盘的孔中。此外,轻轻地排出水,以免干扰密封的花瓣。用一条透明胶带密封注水孔。
  14. 标记圆柱。我们编号(或字母)我们的圆柱体,并记录该样品的描述,包括植物种类,植物部分,表面,样品的年龄,处理和日期。
  15. 将圆筒放入塑料容器中,并返回培养箱。
  16. 对要分析的每个样品重复步骤8-15
  17. 将圆筒放在25℃的培养箱中的干燥剂容器中过夜以稳定
  18. 称量气瓶,记录测量时间和质量,并将它们返回到干燥剂容器和培养箱。虽然是可选的,通过数据记录程序将天平连接到计算机,以Microsoft Excel兼容文件格式记录时间和质量,这简化了数据处理。假设在称量气缸之前,已经根据制造商的说明校准了天平。我们还建议称量与样品质量相似的标准品。这允许在进行几天的测量时确认天平的精度(参见下面的步骤)。
  19. 在设定的时间间隔重复步骤18。经过的时间必须允许足够的水损失,使得新质量从先前质量减少大于天平的误差。对于相当可渗透的植物表面, 宇宙花瓣(Buschhaus ,2015),每2小时测量一次,因为我们的平衡d = 0.1mg。然而,样品通常每天称重一次以减少百分比误差。更多的防水植物表面可能需要比这更长的间隔。时间间隔可以增加或减少以考虑到您的平衡的精度 当每个气瓶被取出进行称重时,检查下面的干燥剂的颜色。如果紧在圆柱体下方的干燥剂变为水合色,则植物圆盘可能破裂。轻轻转动圆筒,检查植物圆盘是否有肉眼可见的撕裂,并丢弃撕裂的样品
  20. 绘制每累积时间变化的质量的累积变化。线性回归线通常占观察到的变化的97%以上。当r 2 值小于0.97时,对图的快速检查通常显示出在植物盘破裂或室耗尽水的点处的线的明显弯曲。代表性数据可以在Buschhaus等人的图2(2015)中找到。
  21. 根据等式P = F A -1 C -1 计算磁导P(m s <-1)。 F是流量(mg s -1 ),等于来自前一步骤的线性回归线的斜率。 A是叶盘的暴露区域。确保这是真实的表面积,而不只是表观表面积。这对于瓣的近轴表面尤其重要,其中表皮细胞通常不是平的。 C是水蒸气浓度的变化(在25℃下23.07mg m -2 -3)。公约要求报告几何平均值,因为数据通常偏向更高的渗透(Baur,2008)。关于方程的进一步解释,参见Schreiber和Sch?nherr(2009)。

    图4.用于水渗透的重量测量的圆柱体。硅脂(白色带)施加在盖子和圆筒底部(A)上的环中。瓣膜盘被放置在盖子和底部(B)中的开口之间,并且盖子被牢固地压在底部(C)上。盖子和底部用一条胶带(阴影白色; D)固定在一起以完成圆筒组件(E)。


  1. 气缸的尺寸和成分是适应性的。为了便于后续计算,进入圆筒中心的开口应具有恒定的直径。我们发现11mm的直径对于宇宙花瓣是令人满意的,并且还允许圆柱体用于分析来自其他物种的叶子。内部容积应至少含有1毫升水,因为这样可以在用完之前测量水分损失(平衡物的质量变化达到1克,读数为0.0001克)。在底部钻有直径为2mm的孔,以便在花瓣被密封穿过开口之后允许圆筒充满水。在我们的情况下,圆柱体由不锈钢加工而成。然而,任何不能渗透水的材料应该是足够的。外部尺寸为约2.5cm×2cm(直径高度)。
  2. 为了测试你的密封技术,密封一个一分钱横跨圆筒的开口,而不是植物圆盘。使用此设置,不应观察到质量的变化。
  3. 虽然对于花瓣(或异常叶片)而言不是问题,但是许多叶片具有提供用于水运动通过屏障的替代路线的气孔。气瓶可在促进气孔闭合的条件下温育,但在解释结果时仍必须小心。
  4. 这个实验设置可以探索几个变化。在植物盘中对屏障进行定量之后,可以移除表皮蜡层并重新分析盘。之后,可以去除所有的表皮蜡并对盘进行再分析。以这种方式,可以确定表皮蜡层的相对贡献。
  5. 最简单的植物生长发生在将宇宙直接接种到土壤上(例如阳光混合物4;每15cm直径的盆中有1棵植物),每周用肥料浇水以及在12℃下恒定20℃温度的生长条件h天(约150μmolm -2光生物活性辐射)/12夜条件。如果空间有限,可以在移植到土壤之前先播种种子,如Buschhaus等人(2015年)所述,以确保每个花盆都有成功生长的植物。


作者感谢L. Schreiber博士(德国波恩大学)建议表面磨损物的构造。在该公开中使用的方法首先在Sch?nherr和Lendzian(1981)中描述,并且首先适应于Buschhaus等人的花瓣。 (2015)。对自然科学和工程研究委员会(加拿大),加拿大研究椅计划和加拿大创新基金会提供了对R. Jetter的大力支持。


  1. Baur,P。(2008)。 植物中水渗透性和有机溶质迁移率的对数正态分布角质层。植物细胞环境20(2):167-177。
  2. Buschhaus,C.,Hager,D。和Jetter,R。(2015)。 宇宙七叶树花瓣上的蜡层对总花瓣水阻力的贡献不等。 植物生理学 167(1):80-88
  3. Schreiber,L。和Sch?nherr,J。(2009)。植物角质层的水和溶质渗透性。 Springer-Verlag Berlin Heidelberg。
  4. Sch?nherr,J。和Lendzian,K。(1981)。 测量孤立的植物角质层膜的透水性的简单而便宜的方法。 Z Pflanzenphysiol 102(4):321-327。
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引用: Readers should cite both the Bio-protocol article and the original research article where this protocol was used:
  1. Jetter, R. and Buschhaus, C. (2015). Quantifying the Permeability of the Apoplastic Water Barrier in Cosmos Petals. Bio-protocol 5(22): e1652. DOI: 10.21769/BioProtoc.1652.
  2. Buschhaus, C., Hager, D. and Jetter, R. (2015). Wax layers on Cosmos bipinnatus petals contribute unequally to total petal water resistance. Plant Physiol 167(1): 80-88.

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