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Molecular resolution atomic force microscopy of soluble proteins in solution
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etBiochi~ic~a Biophysicat~ta
Biochimica et Biophysica Acta 1199 (1994) 105 - 114
Molecular resolution atomic force microscopy of soluble proteins in solution Jie Yang, Jianxun Mou, Zhifeng Shao
*
BIO-SPM Laboratory and Department of Molecular Physiology and Biological Physics, University of Virginia School of Medicine, Box 449, Charlottesville, VA 22908, USA (Received 19 July 1993)
Abstract
We introduce a simple specimen preparatory method for atomic force microscopy of soluble proteins in aqueous solutions. It is demonstrated that the mica surface is suitable for direct adsorption of macromolecules that are sufficientlystable to withstand the disturbance of the probe for reproducible imaging at high resolution. It is also shown that the main problem impeding successful imaging is the excessive adsorption of macromolecules, as loosely bound macromolecules readily stick to the tip and produce various imaging artifacts. Key words: Atomic force microscopy; Ferritin; Apoferritin; Immunoglobulin; Hemoglobin; 1,2-Dihexadecanoyl-rac-glycero-3phosphatidylcholine (DPPC) 1. Introduction
The atomic force microscope (AFM) [1] uses a cantilever mounted force sensor to probe the specimen surface in order to obtain topographical information. With AFM, atomic resolution can be routinely obtained on crystalline materials [2-5]. Since the AFM does not require the specimen to be electrically conductive and can be easily fitted to operate in buffer solutions when optical detection is used to measure the cantilever deflection [6], it should be the preferred choice for biological applications. Indeed, despite our limited understanding of the contrast mechanism of AFM on deformable surfaces, many reproducible high resolution images of biological specimens have been published, such as DNA [7-13], membrane proteins [14-16] and phospholipid bilayers [17,18], among many others [19; for a review, see Refs. 20, 21 and references therein]. Together with the study of dynamics of macromolecules [3,22,23], the power and usefulness of AFM for structural biology has been convincingly demonstrated. However, the performance and application of AFM in biology are still limited by many factors, most of
* Corresponding author. Fax: + 1 8049821616. 0304-4165/94/$07.00 © 1994 Elsevier Science B.V. All rights reserved SSD! 0 3 0 4 - 4 1 6 5 ( 9 3 ) E 0 1 0 2 - O
which we have just started to understand. Among these factors, the softness of most biological materials has been suggested as one of the major limitations for achieving high resolution on most biological samples [20,21]. To solve this problem without extensive manipulation of the macromolecules themselves may require the use of cryogenic temperatures, at which the mechanical strength of most proteins should be significantly improved [24-26]. The instrumental development required for this purpose has just been started [27-29], and may eventually further push AFM resolution of biological samples down to the sub-nm regime. For room temperature imaging, particularly in buffer solutions, it has been argued that the biggest problem was how to 'fix' these macromolecules to the substrate, without the disturbance of the probe damaging or moving them around [30]. For imaging membrane proteins, the newly developed method, using Langmuir trough prepared lipid membranes on a solid support with reconstituted proteins [16], appears to be generally applicable for high resolution imaging; this method has already yielded 1-2 nm resolution on randomly distributed cholera toxin, comparing well with electron microscopy of 2-d crystals [31-33]. But in the case of soluble proteins, the problem is perceived to be much more difficult, since the stabilizing effect of a lipid bilayer is no longer available and one must rely on the
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direct interaction between the macromolecules and the substrate. Considering the relatively large probe force ( ~ sub-nN) and the thermal fluctuation at room temperature, simple adsorption did not appear sufficient for AFM imaging. In fact, such direct adsorption has not yielded any reproducible images at reasonable resolution. To circumvent this problem, a number of schemes were proposed, such as binding the proteins to an artificial molecular layer [34], or modifying the proteins so that they could bind to a specially treated substrate [35]. Unfortunately, these methods are quite material specific and may not be generally applicable to other proteins. An even greater problem is that the molecular structure of proteins may be altered by such modifications. To make AFM a generally useful laboratory instrument, like the electron microscope, a simple and generally applicable specimen preparatory method must be found. In this article, we demonstrate, via the imaging of several soluble proteins at molecular resolution, that the mica surface is suitable for stable adsorption of soluble proteins to allow reproducible AFM imaging in aqueous solutions. The resolution is quite comparable to that of electron microscopy of negatively stained specimens, but is obtained under much more natural conditions. We will show that the major problems of AFM imaging of these soluble proteins are not stable adhesion of the proteins to the substrate. In fact, for most cases, it was found that mica surface was covered with macromolecules and those loosely attached molecules, probably above the first few layers, readily stuck to the tip, producing various false images. The best specimens are those with only a monolayer or a few layers of macromolecules at the surface. Only then, reproducible and stable AFM imaging can be achieved with a very clean tip.
2. Materials and methods
vided in the captions of corresponding pictures) was applied to a mica surface which had already been examined by AFM in solution without proteins, as a control. The protein was allowed to adsorb to the surface in a period of 1/2-5 min and followed by a gentle wash with either deionized water or a low salt buffer (15 mM NaCI, 1 mM phosphate buffer, with or without 1 mM EDTA). The specimen was never allowed to dry and was imaged by AFM in deionized water or a low salt buffer. Method 2: Freshly cleaved mica substrate was immersed in a mini-trough made of teflon, filled with solutions of corresponding proteins. To reduce protein consumption, the mini trough has a small volume of 0.2 ml. The mini-trough was submerged in a water bath to control the incubation temperature. In most cases, the temperature was increased to about 40°C and kept at that temperature for 0.5-1 hour. This was followed by very slow cooling (2-6 hours). Prior to AFM imaging, the specimen was again gently washed with either deionized water or a low salt buffer. 2.3. A F M imaging
A home made fluid cell retrofitted to a NanoScope II AFM (Digital Instruments, Santa Barbara, CA) and Si3N 4 cantilevers (k = 0.06 N / m ) were used for imaging. For specimens prepared by method 1, freshly cleaved mica was imaged in low salt buffer or deionized water prior to application of protein solutions. The mica surface normally appeared fiat and clean, even after storage in deionized water overnight. For minimal specimen deformation or tip damage, initial engagement of the tip was carried out over a small scanning area. After examination of the probe force, it was adjusted to below 0.2 nN and followed by larger area scanning to image the proteins. The typical scanning speed was 4.7 Hz and all images were taken in the constant force mode at room temperature.
2.1. Materials
3. Results and discussion
Ferritin and apoferritin (from horse spleen), Immunoglobulin G (IgG) conjugated with ferritin, Immunoglobulin M (IgM from mouse), Immunoglobulin A (IgA from mouse), synthetic 1,2-Dihexadecanoylrac-glycero-3-phosphatidylcholine (DPPC) and hemoglobin (from pig), were purchased from Sigma Chemical Co. (St. Louis, MO) and used directly without further purification.
In this section, we present results of atomic force microscopy of several soluble proteins to demonstrate that mica is an excellent substrate that provides very stable specimens for AFM imaging in aqueous solutions. We will, with each case, discuss the problems encountered in high resolution AFM imaging and the precautions that must be observed, in order to obtain reproducible images of good quality.
2.2. Specimen preparation 3.1. Ferritin and apoferritin
Method 1: At room temperature, 20 /zl of protein solution (the protein concentrations used were between 5 /zg/ml to 5 m g / m l and the details are pro-
Ferritin is an iron storage protein which plays an important role in metabolism [36,37]. It has an irregu-
J. Yang et al./ Biochimica et Biophysica Acta 1199 (1994) 105-114
larly shaped core of iron-oxide microcrystals with a size of 5 - 7 nm, containing 4000-7000 iron atoms. The protein shell, apoferritin, consists of 24 subunits forming a dodecahedron surrounding the core with a total molecular weight of about 440 kDa [36,37]. Depending on the orientation of the molecule on the substrate, the projection of the molecule can be a square with 11 nm sides, a hexagon with 6.4 nm sides, or a hexagon with four sides of 6.4 nm and two sides of 8.3 nm [38]. In the last case, the molecule should measure about 11 × 14 nm. Fig. 1 shows two typical AFM images of ferritin obtained in deionized water. In Fig. l(a), ferritin molecules are seen closely packed (method 1) and in Fig. l(b), loosely distributed (method 2). In neither case is long range order observed. These specimens were very stable for repeated scans and could be stored at 4°C for several days without noticeable degradation. A probe force of up to 0.8 nN could be applied without visible change of the image. At the resolution already achieved, the molecular orientation can be directly identified in A F M images. In Fig. 2(a), the ferritin molecules are seen predominantly having an elliptical shape, while in Fig. 2(b), they appear spherical. In the latter case, whether they are a square or a hexagon is not resolved due to the limited resolution achieved on these specimens, although the dimension is roughly correct. But with exceptionally good tips, the subunit structure within each ferritin molecule was nearly resolved (Fig. 2(c)), even without correspondence analysis. A comparison with the theoretical model of various molecular orientations is shown in Fig. 2(d), and the consistency is more than apparent. Correspondence analysis of such images might be useful in further interpreting the data [39]. These results compared quite well with those from electron microscopy of negatively stained ferritin specimens, where the subunit structure was not resolved normally. We should also point out that the lateral molecular dimensions, determined from both loosely distributed and closely packed molecules, are surprisingly close to the correct values determined from other techniques (see Table 1) with very small tip-related broadening ( < 1 nm). If one examines a hard ball model (see Fig. 3) for both loosely distributed and closely packed specimens, the tip radius of curvature, Rt, in the absence of any surface deformation, can be calculated from either the half height full width (for loosely distributed molecules) or from the contrast (for closely packed molecules). From the half height full width (D) of individual molecules, we have
Fig. 1. AFM images of ferritin obtained in deionized water with a probe force about 0.1 nN and a scanning speed of 4.73 Hz. (a) shows a closely packed specimen prepared by method 1 (5 mg/ml ferritin in 0.15 M NaC1). (b) is a typical image of a loosely distributed specimen prepared by method 2 where the incubation temperature was 40°C (200 mg/ml ferritin in 0.15 M NaCI). No filtering was applied.
contrast of closely packed molecules, we have (r--Ah) 2 Rt =
(a/2)2-r Rt =
2r
2
(1)
where r is the radius of the hard ball. From the
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2A h
(2)
where A h is the measured height contrast. Based on this model, our data would indicate a tip radius of curvature of about 0.5 nm according to Eq. (1) (for A = 12 nm and 2r = 11 nm), which is too small to be
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trustworthy and about 5 nm according to Eq. (2) (where the measured contrast is about 1.5 nm), which is still much smaller than the value reported elsewhere (30-50 nm) [40]. Particularly, we noticed that the height of the molecules, measured from loosely distributed speci-
mens, is only about 40% (between 4-6 nm) of the correct value, indicating quite noticeable vertical compression even at a probe force of ~ 0.1 nN. Normally, one would expect even lower lateral resolution when deformation is severe. Such an assertion does not seem
i
ii
iii
2 0 nm
d Fig. 2. A F M images of ferritin molecules to show molecular orientation and details. In (a), the molecules appeared elliptical in shape, but in (b), spherical shape is seen, corresponding to a different molecular orientation. Above images do not have sufficient resolution to resolve the subunit structure. But with exceptionally good tips, more details were revealed. Shown in (c) is an example. These structures were reproduced by several tips on the same specimen. A comparison with the model u n d e r different orientations is shown in (d). T h e model is slightly tilted to show the 3-d perspective. It is seen that the details are consistent with the known subunit structure. All images were obtained in deionized water with the probe force in the range of 0.1 x 0.2 n N and the scanning speed of 4.73 x 5.79 Hz. These specimens were prepared by method 1. Low pass filtering was used to reduce high frequency noise.
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Table 1 Lateral dimensions from AFM images Ferritin Apoferritin
Elliptical Spherical Elliptical Spherical
Ferritin-IgG
Length (nm)
Width (nm)
14.5 + 1.0 11.5 + 0.9 15.1:1:1.1 12.0 + 1.0 22 + 3
11.0 :t: 1.0 10.9 :i: 0.9 11.8:1:1.2 11.8 + 1.3 17 + 3
consistent with our data. At present, the relationship between lateral resolution and vertical compression is basically not understood. It may be argued that the exceptional stability and small tip broadening of ferritin molecules might be due to the iron-oxide core that increased the mechanical rigidity of the molecule. To test whether this is essential for stable AFM imaging at high resolution, apoferritin, the protein shell without the iron-oxide core was also imaged by AFM in deionized water. Fig. 4 is a typical image of apoferritin prepared by method 2. We found that apoferritin was a little less stable than complete ferritin, but incubation at 40°C improved the specimen stability significantly. The reason for the improvement is not clear. It is seen that the molecular dimensions are slightly larger than that of complete ferritin molecules (Table 1). This result indicates that the iron-oxide core makes little contribution to specimen stability or mechanical strength, because, surpisingly, even the molecular height is very close to that of the complete ferritin. No central depression was ever observed, contrary to what one would expect, suggesting that the protein shell itself is as strong as the complete ferritin. These results demonstrate that, with-
(a)
~
~
'~--'
' (b)
]
2r
~
Rt Ah
Fig. 3. Hard ball model to illustrate the tip broadening effect. The end of the tip is represented by a sphere of radius R t and related parameters for Eq. (1) and Eq. (2) are shown correspondingly. In (a), an isolated molecule is shown where the half height full width A is normally measured. In (b), a closely packed case is shown where the contrast d h can be measured.
Fig. 4. A typical apoferritin image obtained in deionized water at a probe force of ~ 0.1 nN and a scanning speed of 4.73 Hz. Notice the similarity between this image and that of ferritin molecules. The specimen was prepared by method 1 with the protein solution containing 5 m g / m l apoferritin and 0.15 M NaC1. No filtering was applied.
out modification of the substrate (mica) or the protein themselves, the AFM is capable of high resolution imaging of soluble proteins at room temperature in aqueous solutions. 3.2. Ferritin-IgG conjugates
Although specimens of ferritin and apoferritin prepared by both methods showed excellent stability on mica surface in solution, initial application of the method to ferritin-IgG conjugates was difficult and confusing. Unlike the case of imaging ferritin and apoferritin, where the adhesion force between the tip and the specimen was not detected, it was found that with ferritin-IgG conjugates, an unstable adhesion force often appeared, up to 10 nN in some cases. In the presence of such adhesion forces, the AFM images obtained were often rather flat without much feature, except a few large aggregates that showed noticeable drift with repeated scans, perhaps being pushed around by the tip. However, with a different tip when the adhesion force was absent, the same specimen could show good coverage of molecules on the surface, although other imaging conditions were almost identical. We found that, in some cases, particularly with those specimens prepared by method 2, the adhesion force can actually be very small ( < 2 nN) or absent. Only in these cases were stable images of ferritin-IgG conjugates obtained (see Fig. 5). In Fig. 5(a), ferritin-IgG
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J. Yang et al. /Biochimica et Biophysica Acta 1199 (1994) 105-114 ally polymerize under certain conditions seems to indicate that the preferred IgG binding sites have certain symmetry that can facilitate inter-molecular interactions that favor some long range order. We should also point out that electron microscopy also failed to differentiate IgG molecules from ferritin unequivocally in ferritin-IgG conjugates [41]. The existence of large adhesion force in solution on these specimens is not fully understood at present. However, a plausible explanation is that loosely attached molecules (beyond the first few layers) became associated with the tip after engagement. This explanation would be consistent with the case of imaging D N A in air, where the adhesion force has been attributed to tip contamination [8,12]. Unfortunately, cleaning the tip on carbon coated mica did not yield consistent results for ferritin-IgG specimens. But we did observe that for tips having a large adhesion force, vigorous washing with salt solutions (PBS and 1 mM EDTA, or 1 M NaC1) often yielded some improvement on both the adhesion force and the image quality, probably due to the removal of some of the contaminants by the solution. These results indicate that the condition of the tip is still the most important factor in AFM imaging of biological samples and is also the most difficult to control. 3.3. Immunoglobulin M (IgM)
Fig. 5. Images of ferritin-IgG conjugates with tips showing no adhesion force. In (a), ferritin-IgG conjugates are seen polymerized.This image was obtained in 12 mM NaCI, 0.3 mM KCI and 1 mM phosphate buffer, pH 7.0. In (b), loosely distributed ferritin-IgG conjugates were imaged in deionized water. The irregular shape and the larger size are in clear distinction from those images of ferritin and apoferritin (Fig. 1 and Fig. 4). Scale bars: 200 nm (horizontal) and 50 nm (vertical). Both samples were prepared by method 2 with 100 ~g/ml ferritin-IgG conjugates in PBS buffer. No filtering was applied.
molecules are seen to polymerize (for unknown reasons) on the mica surface and in Fig. 5(b), lower coverage revealed individual molecules, although they were rarely completely isolated. According to Sigma Chemicals, each ferritin should have, on average, 2.3 IgG molecules bound. But the A F M did not resolve each IgG molecules from the ferritin, probably due to the flexibility of the IgG and the shadowing effect of the ferritin (since the ferritin should be much more rigid with a larger height than the IgG). But the molecular profile is considerably different from both ferritin and apoferritin. The measured lateral size of ferritin-IgG conjugate is about 22 nm × 17 nm, consistent with the assumption of one ferritin with two IgG bound. The fact that ferritin-IgG conjugate can actu-
In view of the difficulty with ferritin-IgG conjugates, the presence of IgG molecules on the surface of ferritin apparently complicated the tip-sample interaction. Thus, we further tested the method with a pure antibody specimen - IgM. IgM consists of 5 IgG subunits and a J chain at its center, connected through disulfide bonds between neighboring IgGs and between the IgG and the J chain. The total molecular weight is between 900 and 1000 kDa [42]. The only direct structural elucidation of IgM is from electron microscopy which showed 5 dendrite like spikes radiating from the center [41,43]. The IgM proved to be even more difficult to image than ferritin-IgG conjugates. Although IgM readily adhered to the mica surface under various conditions, the tip-sample interaction is extremely complicated. The most notable phenomenon is the appearance of negative (or so-called inverted) contrast. Fig. 6 shows an example in which all molecules showed a negative contrast, although the specimen was quite stable with repeated scans of good reproducibility under the same tip. It was observed that for a tip that yielded negative contrast, the image occasionally reverted to positive contrast, but in this case stable imaging was often impossible. We believe that the negative contrast is the direct consequence of tip contamination, which was always accompanied by a measurable adhesion force.
J. Yang et aL / Biochimica et Biophysica Acta 1199 (1994) 105-I14
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Fig. 6. A negative contrast image of IgM, as observed when the tip
2 5 0 nm
had an adhesion force larger than 2 nN. The size of individual molecules is that of IgM (35 + 5 nm), but none of them showed any detail. With different tips/specimens, the molecular size of negative contrast images varied by a factor up to 2. This specimen was prepared by method 2 with an incubation temperature of 40°C (50 mg/mi IgM in PBS buffer). No filtering was applied.
Fig. 7. A typical IgM image obtained in deionized water at a scanning speed of 4.73 Hz and a probe force of 0.2 nN without adhesion force. The specimen was prepared by method 2 at an incubation temperature of ~ 45°C (20 m g / m l IgM in PBS). The specimen showed good protein coverage, with an average molecular size of 35 + 3 nm. The same specimen could show negative contrast for some tips with non-zero adhesion force. See text for more discussion. No filtering was applied.
For good tips with very small adhesion force, positive contrast images could be obtained on the same specimen that yielded negative contrast for other tips. Fig. 7 is a typical 'normal' image of IgM in deionized water as prepared by method 2. Fig. 8 shows a gallery of 10 typical IgM molecules selected from several spec-
imens. It is seen that both the five IgG subunits and the central ring are resolved. The variation in morphology is expected from such large flexible molecules. The
a
b
c
d
e
f
g
h
i
j
50 nm Fig. 8. A collection of 10 typical IgM images. The 5 IgG subunits are clearly recognizable, with finer details revealed in some of them. The variation in morphology may be the result of the flexibility of the molecules in solution which is consistent with electron microscopy results. Low pass filtering was applied to reduce high frequency noise.
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quality of these images are quite comparable to that of electron microscopy [41,43]. Measurements showed a molecular diameter of 35 + 3 nm, which is 17% larger than the size determined from electron microscopy, perhaps due to the incubation at 45°C which broke some of the disulfide bonds, resulting in a more extended molecular profile upon adsorption, although we cannot rule out the possibility that some tip broadening is involved. A n o t h e r possibility is that the dimension from electron microscopy is an underestimate due to possible shrinkage in negative staining [44]. Since no further purification was p e r f o r m e d , the smaller molecules seen in Fig. 7 could be the contaminants or the debris of IgM broken apart during incubation. The height measured from the A F M images is between 2 - 5 nm, which showed a relatively smaller compression. It is interesting to note that the height variation due to the probe force was also quite small for D N A and bacteriophage adsorbed on aminopropylytriethoxy silane treated mica when imaged in water. The reason for this p h e n o m e n o n is not understood [8]. One would normally expect the IgM to be more compressible than the ferritin. These images were reproducible and stable with fresh tips that showed positive contrast without adhesion force. These specimens could be stored at 4°C for more than 48 h. The A F M results of IgM once again indicated the importance of controlling the tip condition. An additional point raised from this experiment is that one must be careful in interpreting the A F M images, due to the existence of negative contrast. This is particularly a problem for closely packed specimens, where false results can easily arise from the negative contrast that may be difficult to recognize.
a
b
3 0 nm
3.4. Applications to several other macromolecules
Fig. 9. AFM images of IgA molecules obtained in deionized water with a probe force of 0.1 nN and a scanning speed of 4.73 Hz. Specimens were prepared by method 2 at an incubation temperature of 40°C (Protein concentration: 5 /zg/ml in PBS). Images shown were obtained after overnight storage at 4°C. (a) shows a large scale image where IgA molecules apparently polymerized.The double line structure should be the reflection of the IgG dimer in leA. The arrow indicates one such dimer that can be directly identified. (b) shows four examples of individual IgA molecules. Each monomer measures about 17 nmX 11 nm, consistent with the IgG dimension. Further details were not resolved at this resolution. Low pass filtering was applied to reduce high frequency noise.
To make sure that the method is indeed applicable for routine experiments, we have applied the method outlined in Sec. II to few m o r e macromolecules. Fig. 9 is the A F M image of I g A obtained in deionized water. The specimens were p r e p a r e d by method 2. IgA has been known to be a dimer of I g G molecules linked through a J chain [45]. Interestingly, IgA also polymerized on the mica surface (Fig. 9(a)), like the ferritin-IgG conjugates (Fig. 5(a)). However, the close association of IgA molecules makes it difficult for direct identification of each dimer in the image. F r o m these polymers, the I g A molecule has a normal size of 24 + 3 nm x 17 + 2 nm, consistent with the known values based on the I g G structure. On some areas, though, the dimer configuration could be clearly identified. Fig. 9(b) shows four such images. It is seen that each m o n o m e r has a size of ~ 17 x 11 nm and the relative position is somewhat variable, as one would expect. These values are consistent with an orientation in which the IgA
molecule is fully extended. Fig. 10 is an A F M image of phospholipid vesicles (DPPC) adsorbed on mica surface, p r e p a r e d by method 1. The vesicles were prepared by the method outlined by H u a n g [46], which should be primarily unilamella vesicles. At room temperature, D P P C should be in the gel phase (transition temperature: 41°C). These vesicles were surprisingly stable, with almost no adhesion force. If one would attribute protein adhesion to mica as electrostatic, since mica is negatively charged, these vesicles would not fit into the explanation, because phosphatidylcholine is neutral. One thing to notice is that the top of these vesicles was not flattened by the probe force, contrary to one might expect for these relatively large holo-structures (the average diameter of these vesicles is 48 nm, but the largest one has a diameter of 200 nm). This result seems in contradiction to that of ferritin (Fig. 1). Further study is needed to elucidated
J. Yang et al. /Biochimica et Biophysica Acta 1199 (1994) 105-114
4/ Fig. 10. This is an AFM image of DPPC vesicles adsorbed on mica surface. The majority of the vesicles are smaller ones and their average diameter is 485:7 nm. The vesicles were prepared by the method described by Huang [46] with a buffer solution containing 140 mM NaCI, 10 mM Tris, pH 8.0. The specimen was prepared by method 1. The image was obtained in the same buffer with a probe force of 0.5 nN and a scanning speed of 4.73 Hz. It is noticed that the top of the vesicles, even the larger ones, is not flattened by the probe. No filtering was applied. Scale bars: 500 nm (horizontal) and 180 nm (vertical).
this phenomenon. If proteins are incorporated in these vesicles, it is highly likely that they can be resolved by AFM. Fig. 11 is the AFM image of hemoglobin, taken in deionized water. The specimen was prepared by method 2, but incubated at 4°C overnight for near monolayer coverage. With hemoglobin, we found that the proteins stuck to the tip very easily when the protein coverage on the substrate was more than one layer. For those good specimens, the image was very stable and reproducible, as long as the probe force was kept below 0.2 nN. The molecular dimension measured
Fig. 11. AFM image of hemoglobin on mica surface. The specimen was prepared by method 2 (5 /Lg/ml hemoglobin in PBS) incubated at 4°C overnight. The hemoglobin measures 6.5-I-0.9 nm, slightly larger than the result from X-ray diffraction [47], maybe an indication of some tip-related broadening. This image was obtained in deionized water with a probe force of ~ 0.1 nN and a scanning speed of 4.73 Hz. Low pass filtering was applied to reduce high frequency noise. Scale bars: 50 nm (horizontal) and 12 nm (vertical).
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is 6.5 + 0.9 nm, slightly larger than the actual size (5.0 nm × 5.5 nm × 6.4 nm) determined from X-ray diffraction [47]. These additional AFM images of various macromolecules convincingly demonstrated that the methods presented in Section 2 are indeed generally applicable to soluble macromolecules and mica surface is adequate for nm resolution atomic force microscopy of directly adsorbed macromolecules in aqueous solutions.
4. Concluding remarks Specimen preparation is perhaps the most important aspect of any imaging technique. This is particularly true for AFM, which was invented only a short few years ago. Only recently, the AFM for biological applications began to appear promising, as represented by the success with DNA and membrane proteins. In this study, it is demonstrated that, contrary to popular perception, mica is a remarkable substrate not only for membranes, but also for soluble proteins. The method of direct adsorption on mica surface yielded stable, reproducible and high resolution AFM images at nearly physiological conditions. Our data also indicated that the gross conformation of macromolecules was little changed by the surface interaction with mica, at least, at the resolution achieved. We are surprised at the finding that, the main problem of AFM of these soluble proteins is not the adhesion to the substrate. In fact, all soluble proteins we have tried, including albumin, IgG, hemocyanin and myosin, adhered well to the substrate and often too much rather than too little. The problem is then how to prepare a monolayer or just a few layers on a substrate (not necessarily mica) with good uniformity. An additional but very serious problem is how to prevent proteins from sticking to the tip, a factor we have very little control of at present. This may require the use of different tip materials. If indeed these materials can be found, we can then further experiment with the tip geometry in order to improve upon the resolution achieved so far. There is reason to believe that with a good monolayer coverage and a non-sticking tip, similar resolution as the cholera toxin (1-2 nm) should be achievable by this method. A minor discovery from these experiments is that many proteins polymerized upon adsorption on mica surface for unknown reasons. In principle, this could provide some indirect information about the conformation of these macromolecules. Considering the short history of biological AFM, with further refinement of both specimen preparation techniques and tip fabrication, the AFM should become a useful imaging instrument to supplement with other techniques. The AFM should be able to provide
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a resolution comparable to that of electron microscopy with a specimen preparation method not much more complicated than that for light microscopy.
Acknowledgements We would like to thank Drs. A.P. Somlyo, A.V. Somlyo and S.M. Lindsay for useful discussions. We would also like to thank Zhiyu Hu for technical assistance. Support from Whitaker Foundation, US Army Research Office, Jeffress Memorial Trust, National Science Foundation and National Institutes of Health is gratefully acknowledged.
References [1] Binnig, G., Quate, C.F. and Gerber, Ch. (1986) Phys. Rev. Lett. 56, 930-933. [2] Albrecht, T.R. and Quate, C.F. (1987) J. Appl. Phys. 62, 25992602. [3] Drake, B., Prater, C,B., Weisenhorn, A.L., Gould, S.A.C., Albrecht, T.R., Quate, C.F,, Cannell, D.S., Hansma, H.G. and Hansma, P.K. (1989). Science 243, 1586-1589. [4] Gould, S.A.C., Burke, K. and Hansma, P.K. (1989) Phys. Rev. B40, 5363-5366. [5] Ohnesorge, F. and Binnig, G. (1993) Science 260, 1451-1456. [6] Alexander S., Hellemans, L., Marti, O. Schneir, J. Elings, V. and Hansma, P.K. (1989) J. Appl. Phys. 65, 164-167. [7] Lyubchenko, Y.L., Jacobs, B.L. and Lindsay, S.M. (1992) Nucleic Acids Res. 20, 3983-3986. [8] Lyubchenko, Y.L., Oden, P.I., Lampner, D., Lindsay S.M. and Dunker, K.A. (1993) Nucl. Acids Res. 21, 1117-1123. [9] Bustamante, C., Vesenka, J., Tang, C.L., Rees, W., Guthod M. and Keller, R. (1992) Biochemistry 31, 22-26. [10] Yang, J., Takeyasu K. and Shao, Z. (1992) FEBS Lett. 301, 173-176. [11] Hansma, H.G., Vesenka, J., Siegerist, C., Kelderman, G., Morrett, H., Sinsheimer, P.L., Elings, V., Bustamante C. and Hansma, P.K. (1992) Science 256, 1180-1184. [12] Yang, J. and Shao, Z. (1993) Ultramicroscopy, in press. [13] Rees, W.A., Keller, R.W., Vesenka, J., Yang, G. and Bustamante, C. (1993) Science 260, 1646-1649. [14] Butt, H.-J., Downing, K.H. and Hansma, P.K. (1990) Biophys. J. 58, 1473-1480. [15] Hoh, J.H., Lal, R., John, S.A., Revel, J.-P. and Arnsdorf, M.F. (1991) Science 253, 1405-1408. [16] Yang, J., Tamm, L.K., Tillack, T.W. and Shao, Z. (1993) J. Mol. Biol. 229, 286-290. [17] Hansma, H.G., Gould, S.A.C., Hansma, P.K., Gaub, H.E.,
Longo, M.L. and Zasadzinski, J.A.N. (1991) Langmuir 7, 10511054. [18] Zasadzinski, J.A.N., Helm, C.A., Longo, M.L., Weisenhorn, A.L., Gould, S.A.C. and Hansma, P.K. (1991). Biophys. J. 59, 755-760. [19] Hoh, J.H. and Hansma, P.K. (1992) Trends in Cell Biology 2, 208-213. [20] Engel, A. (1991) Annu. Rev. Biophys. Biophys. Chem. 20, 79108. [21] Yang, J., Tamm, L.K., Somlyo, A.P. and Shao, Z. (1993) J. Microscopy, in press. [22] Haberle, W., Horber, J.K.H., Ohnesorge, F., Smith, D.P.E. and Binnig, G. (1992) Ultramicroscopy 42-44, 1161-1167. [23] Henderson, E., Hayelon P.G. and Sakaguchi, D.S. (1992) Science 257, 1944-1946. [24] Prater, C.B., Wilson, M.R., Garnaes, J., Massie, J., Elings, V.B. and Hansma, P.K. (1991) J. Vac. Sci. Technol. B9, 989-991. [25] Rasmussen, B.F., Stock, A.M., Ringe, D. and Petsko, G.A. (1992) Nature 357, 423-424. [26] Perutz, M.F. (1992) Nature 358, 548. [27] Giessibl, F.J., Gerber Ch. and Binnig, G. (1991) J. Vac. Sci. Technol. B9, 984-988. [28] Giessibl F.J. and Binnig, G. (1992) Ultramicroscopy 42-44, 281-289. [29] Mou, J., Yang, J. and Shao, Z. (1993) Rev. Sci. Instrum. 64, 1483- 1488. [30] Keller, D. and Bustamante, C. (1993) Biophys. J, 64, 896-897. [31] Reed, R.A., Mattai, J. and Shipley, G.G. (1987) Biochemistry 26, 824-832. [32] Ribi, H.O., Ludwig, D.S., Mercer, K.L., Schoolnik, G.K. and Kornberg, R.D. (1988). Science 239, 1272-1276. [33] Mosser, G. and Brisson, A. (1991) J. Electron Micro. Tech. 18, 387-394. [34] Ohnishi, S., Hara, M., Furuno, T. and Sasabe, H. (1992) Biophys. J. 63, 1425-1431. [35] I11,C.R., Keivens, V.M., Hale, J.E., Nakamura, K.K., Jue, R.A., Cheng, S., Melcher, E.D., Drake, B. and Smith, M.C. (1993) Biophys. J. 64, 919-924. [36] Theil, E.C. (1987) Ann. Rev, Biochem. 56, 289-315. [37] Harrison P.M., Banyard, S.H., Hoare, R.J., Russell, S.M. and Treffry, A. (1977) Ciba Foundation Symposium 51, 19-67. [38] Smith, J.M.A., Stansfield, R.F.D., Ford, G.C. White, J.L. and Harrison, P.M. (1988) J. Chem. Education 65, 1083-1084. [39] Wagenknecht, T., Grassucci, R. and Frank, J. (1988) J. Mol. Biol. 199, 137-147. [40] Albrecht, T.R., Akamine, S., Carver, T.E. and Quate, C.F. (1990) J, Vac. Sci. Technol. A8, 3386-3396. [41] Green, N.M. (1969) Adv. Immunol. 11, 1-30. [42] Metzger, H. (1970) Ann. Rev. Biochem. 39, 889-928. [43] Parkhouse, R.M.E., Askonas, B.R. and Dourmashkin, R.R. (1970) Immunology 18, 575-584. [44] Stokes, D.L. and Green, N.M. (1990) Biophys. J. 57, 1-14. [45] Nezlin, R.S. (1977) Structure and Biosynthesis of Antibodies Consultants. Bureau, New York. [46] Huang, C.-h. (1969) Biochemistry 8, 344-352. [47] Perutz, M.F., Rossmann, M.G., Cullis, A.F., Muirhead, H., Will, G. and North, A.C.T. (1960) Nature 185, 416-422.
ELSEVIER
etBiochi~ic~a Biophysicat~ta
Biochimica et Biophysica Acta 1199 (1994) 105 - 114
Molecular resolution atomic force microscopy of soluble proteins in solution Jie Yang, Jianxun Mou, Zhifeng Shao
*
BIO-SPM Laboratory and Department of Molecular Physiology and Biological Physics, University of Virginia School of Medicine, Box 449, Charlottesville, VA 22908, USA (Received 19 July 1993)
Abstract
We introduce a simple specimen preparatory method for atomic force microscopy of soluble proteins in aqueous solutions. It is demonstrated that the mica surface is suitable for direct adsorption of macromolecules that are sufficientlystable to withstand the disturbance of the probe for reproducible imaging at high resolution. It is also shown that the main problem impeding successful imaging is the excessive adsorption of macromolecules, as loosely bound macromolecules readily stick to the tip and produce various imaging artifacts. Key words: Atomic force microscopy; Ferritin; Apoferritin; Immunoglobulin; Hemoglobin; 1,2-Dihexadecanoyl-rac-glycero-3phosphatidylcholine (DPPC) 1. Introduction
The atomic force microscope (AFM) [1] uses a cantilever mounted force sensor to probe the specimen surface in order to obtain topographical information. With AFM, atomic resolution can be routinely obtained on crystalline materials [2-5]. Since the AFM does not require the specimen to be electrically conductive and can be easily fitted to operate in buffer solutions when optical detection is used to measure the cantilever deflection [6], it should be the preferred choice for biological applications. Indeed, despite our limited understanding of the contrast mechanism of AFM on deformable surfaces, many reproducible high resolution images of biological specimens have been published, such as DNA [7-13], membrane proteins [14-16] and phospholipid bilayers [17,18], among many others [19; for a review, see Refs. 20, 21 and references therein]. Together with the study of dynamics of macromolecules [3,22,23], the power and usefulness of AFM for structural biology has been convincingly demonstrated. However, the performance and application of AFM in biology are still limited by many factors, most of
* Corresponding author. Fax: + 1 8049821616. 0304-4165/94/$07.00 © 1994 Elsevier Science B.V. All rights reserved SSD! 0 3 0 4 - 4 1 6 5 ( 9 3 ) E 0 1 0 2 - O
which we have just started to understand. Among these factors, the softness of most biological materials has been suggested as one of the major limitations for achieving high resolution on most biological samples [20,21]. To solve this problem without extensive manipulation of the macromolecules themselves may require the use of cryogenic temperatures, at which the mechanical strength of most proteins should be significantly improved [24-26]. The instrumental development required for this purpose has just been started [27-29], and may eventually further push AFM resolution of biological samples down to the sub-nm regime. For room temperature imaging, particularly in buffer solutions, it has been argued that the biggest problem was how to 'fix' these macromolecules to the substrate, without the disturbance of the probe damaging or moving them around [30]. For imaging membrane proteins, the newly developed method, using Langmuir trough prepared lipid membranes on a solid support with reconstituted proteins [16], appears to be generally applicable for high resolution imaging; this method has already yielded 1-2 nm resolution on randomly distributed cholera toxin, comparing well with electron microscopy of 2-d crystals [31-33]. But in the case of soluble proteins, the problem is perceived to be much more difficult, since the stabilizing effect of a lipid bilayer is no longer available and one must rely on the
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direct interaction between the macromolecules and the substrate. Considering the relatively large probe force ( ~ sub-nN) and the thermal fluctuation at room temperature, simple adsorption did not appear sufficient for AFM imaging. In fact, such direct adsorption has not yielded any reproducible images at reasonable resolution. To circumvent this problem, a number of schemes were proposed, such as binding the proteins to an artificial molecular layer [34], or modifying the proteins so that they could bind to a specially treated substrate [35]. Unfortunately, these methods are quite material specific and may not be generally applicable to other proteins. An even greater problem is that the molecular structure of proteins may be altered by such modifications. To make AFM a generally useful laboratory instrument, like the electron microscope, a simple and generally applicable specimen preparatory method must be found. In this article, we demonstrate, via the imaging of several soluble proteins at molecular resolution, that the mica surface is suitable for stable adsorption of soluble proteins to allow reproducible AFM imaging in aqueous solutions. The resolution is quite comparable to that of electron microscopy of negatively stained specimens, but is obtained under much more natural conditions. We will show that the major problems of AFM imaging of these soluble proteins are not stable adhesion of the proteins to the substrate. In fact, for most cases, it was found that mica surface was covered with macromolecules and those loosely attached molecules, probably above the first few layers, readily stuck to the tip, producing various false images. The best specimens are those with only a monolayer or a few layers of macromolecules at the surface. Only then, reproducible and stable AFM imaging can be achieved with a very clean tip.
2. Materials and methods
vided in the captions of corresponding pictures) was applied to a mica surface which had already been examined by AFM in solution without proteins, as a control. The protein was allowed to adsorb to the surface in a period of 1/2-5 min and followed by a gentle wash with either deionized water or a low salt buffer (15 mM NaCI, 1 mM phosphate buffer, with or without 1 mM EDTA). The specimen was never allowed to dry and was imaged by AFM in deionized water or a low salt buffer. Method 2: Freshly cleaved mica substrate was immersed in a mini-trough made of teflon, filled with solutions of corresponding proteins. To reduce protein consumption, the mini trough has a small volume of 0.2 ml. The mini-trough was submerged in a water bath to control the incubation temperature. In most cases, the temperature was increased to about 40°C and kept at that temperature for 0.5-1 hour. This was followed by very slow cooling (2-6 hours). Prior to AFM imaging, the specimen was again gently washed with either deionized water or a low salt buffer. 2.3. A F M imaging
A home made fluid cell retrofitted to a NanoScope II AFM (Digital Instruments, Santa Barbara, CA) and Si3N 4 cantilevers (k = 0.06 N / m ) were used for imaging. For specimens prepared by method 1, freshly cleaved mica was imaged in low salt buffer or deionized water prior to application of protein solutions. The mica surface normally appeared fiat and clean, even after storage in deionized water overnight. For minimal specimen deformation or tip damage, initial engagement of the tip was carried out over a small scanning area. After examination of the probe force, it was adjusted to below 0.2 nN and followed by larger area scanning to image the proteins. The typical scanning speed was 4.7 Hz and all images were taken in the constant force mode at room temperature.
2.1. Materials
3. Results and discussion
Ferritin and apoferritin (from horse spleen), Immunoglobulin G (IgG) conjugated with ferritin, Immunoglobulin M (IgM from mouse), Immunoglobulin A (IgA from mouse), synthetic 1,2-Dihexadecanoylrac-glycero-3-phosphatidylcholine (DPPC) and hemoglobin (from pig), were purchased from Sigma Chemical Co. (St. Louis, MO) and used directly without further purification.
In this section, we present results of atomic force microscopy of several soluble proteins to demonstrate that mica is an excellent substrate that provides very stable specimens for AFM imaging in aqueous solutions. We will, with each case, discuss the problems encountered in high resolution AFM imaging and the precautions that must be observed, in order to obtain reproducible images of good quality.
2.2. Specimen preparation 3.1. Ferritin and apoferritin
Method 1: At room temperature, 20 /zl of protein solution (the protein concentrations used were between 5 /zg/ml to 5 m g / m l and the details are pro-
Ferritin is an iron storage protein which plays an important role in metabolism [36,37]. It has an irregu-
J. Yang et al./ Biochimica et Biophysica Acta 1199 (1994) 105-114
larly shaped core of iron-oxide microcrystals with a size of 5 - 7 nm, containing 4000-7000 iron atoms. The protein shell, apoferritin, consists of 24 subunits forming a dodecahedron surrounding the core with a total molecular weight of about 440 kDa [36,37]. Depending on the orientation of the molecule on the substrate, the projection of the molecule can be a square with 11 nm sides, a hexagon with 6.4 nm sides, or a hexagon with four sides of 6.4 nm and two sides of 8.3 nm [38]. In the last case, the molecule should measure about 11 × 14 nm. Fig. 1 shows two typical AFM images of ferritin obtained in deionized water. In Fig. l(a), ferritin molecules are seen closely packed (method 1) and in Fig. l(b), loosely distributed (method 2). In neither case is long range order observed. These specimens were very stable for repeated scans and could be stored at 4°C for several days without noticeable degradation. A probe force of up to 0.8 nN could be applied without visible change of the image. At the resolution already achieved, the molecular orientation can be directly identified in A F M images. In Fig. 2(a), the ferritin molecules are seen predominantly having an elliptical shape, while in Fig. 2(b), they appear spherical. In the latter case, whether they are a square or a hexagon is not resolved due to the limited resolution achieved on these specimens, although the dimension is roughly correct. But with exceptionally good tips, the subunit structure within each ferritin molecule was nearly resolved (Fig. 2(c)), even without correspondence analysis. A comparison with the theoretical model of various molecular orientations is shown in Fig. 2(d), and the consistency is more than apparent. Correspondence analysis of such images might be useful in further interpreting the data [39]. These results compared quite well with those from electron microscopy of negatively stained ferritin specimens, where the subunit structure was not resolved normally. We should also point out that the lateral molecular dimensions, determined from both loosely distributed and closely packed molecules, are surprisingly close to the correct values determined from other techniques (see Table 1) with very small tip-related broadening ( < 1 nm). If one examines a hard ball model (see Fig. 3) for both loosely distributed and closely packed specimens, the tip radius of curvature, Rt, in the absence of any surface deformation, can be calculated from either the half height full width (for loosely distributed molecules) or from the contrast (for closely packed molecules). From the half height full width (D) of individual molecules, we have
Fig. 1. AFM images of ferritin obtained in deionized water with a probe force about 0.1 nN and a scanning speed of 4.73 Hz. (a) shows a closely packed specimen prepared by method 1 (5 mg/ml ferritin in 0.15 M NaC1). (b) is a typical image of a loosely distributed specimen prepared by method 2 where the incubation temperature was 40°C (200 mg/ml ferritin in 0.15 M NaCI). No filtering was applied.
contrast of closely packed molecules, we have (r--Ah) 2 Rt =
(a/2)2-r Rt =
2r
2
(1)
where r is the radius of the hard ball. From the
107
2A h
(2)
where A h is the measured height contrast. Based on this model, our data would indicate a tip radius of curvature of about 0.5 nm according to Eq. (1) (for A = 12 nm and 2r = 11 nm), which is too small to be
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trustworthy and about 5 nm according to Eq. (2) (where the measured contrast is about 1.5 nm), which is still much smaller than the value reported elsewhere (30-50 nm) [40]. Particularly, we noticed that the height of the molecules, measured from loosely distributed speci-
mens, is only about 40% (between 4-6 nm) of the correct value, indicating quite noticeable vertical compression even at a probe force of ~ 0.1 nN. Normally, one would expect even lower lateral resolution when deformation is severe. Such an assertion does not seem
i
ii
iii
2 0 nm
d Fig. 2. A F M images of ferritin molecules to show molecular orientation and details. In (a), the molecules appeared elliptical in shape, but in (b), spherical shape is seen, corresponding to a different molecular orientation. Above images do not have sufficient resolution to resolve the subunit structure. But with exceptionally good tips, more details were revealed. Shown in (c) is an example. These structures were reproduced by several tips on the same specimen. A comparison with the model u n d e r different orientations is shown in (d). T h e model is slightly tilted to show the 3-d perspective. It is seen that the details are consistent with the known subunit structure. All images were obtained in deionized water with the probe force in the range of 0.1 x 0.2 n N and the scanning speed of 4.73 x 5.79 Hz. These specimens were prepared by method 1. Low pass filtering was used to reduce high frequency noise.
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Table 1 Lateral dimensions from AFM images Ferritin Apoferritin
Elliptical Spherical Elliptical Spherical
Ferritin-IgG
Length (nm)
Width (nm)
14.5 + 1.0 11.5 + 0.9 15.1:1:1.1 12.0 + 1.0 22 + 3
11.0 :t: 1.0 10.9 :i: 0.9 11.8:1:1.2 11.8 + 1.3 17 + 3
consistent with our data. At present, the relationship between lateral resolution and vertical compression is basically not understood. It may be argued that the exceptional stability and small tip broadening of ferritin molecules might be due to the iron-oxide core that increased the mechanical rigidity of the molecule. To test whether this is essential for stable AFM imaging at high resolution, apoferritin, the protein shell without the iron-oxide core was also imaged by AFM in deionized water. Fig. 4 is a typical image of apoferritin prepared by method 2. We found that apoferritin was a little less stable than complete ferritin, but incubation at 40°C improved the specimen stability significantly. The reason for the improvement is not clear. It is seen that the molecular dimensions are slightly larger than that of complete ferritin molecules (Table 1). This result indicates that the iron-oxide core makes little contribution to specimen stability or mechanical strength, because, surpisingly, even the molecular height is very close to that of the complete ferritin. No central depression was ever observed, contrary to what one would expect, suggesting that the protein shell itself is as strong as the complete ferritin. These results demonstrate that, with-
(a)
~
~
'~--'
' (b)
]
2r
~
Rt Ah
Fig. 3. Hard ball model to illustrate the tip broadening effect. The end of the tip is represented by a sphere of radius R t and related parameters for Eq. (1) and Eq. (2) are shown correspondingly. In (a), an isolated molecule is shown where the half height full width A is normally measured. In (b), a closely packed case is shown where the contrast d h can be measured.
Fig. 4. A typical apoferritin image obtained in deionized water at a probe force of ~ 0.1 nN and a scanning speed of 4.73 Hz. Notice the similarity between this image and that of ferritin molecules. The specimen was prepared by method 1 with the protein solution containing 5 m g / m l apoferritin and 0.15 M NaC1. No filtering was applied.
out modification of the substrate (mica) or the protein themselves, the AFM is capable of high resolution imaging of soluble proteins at room temperature in aqueous solutions. 3.2. Ferritin-IgG conjugates
Although specimens of ferritin and apoferritin prepared by both methods showed excellent stability on mica surface in solution, initial application of the method to ferritin-IgG conjugates was difficult and confusing. Unlike the case of imaging ferritin and apoferritin, where the adhesion force between the tip and the specimen was not detected, it was found that with ferritin-IgG conjugates, an unstable adhesion force often appeared, up to 10 nN in some cases. In the presence of such adhesion forces, the AFM images obtained were often rather flat without much feature, except a few large aggregates that showed noticeable drift with repeated scans, perhaps being pushed around by the tip. However, with a different tip when the adhesion force was absent, the same specimen could show good coverage of molecules on the surface, although other imaging conditions were almost identical. We found that, in some cases, particularly with those specimens prepared by method 2, the adhesion force can actually be very small ( < 2 nN) or absent. Only in these cases were stable images of ferritin-IgG conjugates obtained (see Fig. 5). In Fig. 5(a), ferritin-IgG
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J. Yang et al. /Biochimica et Biophysica Acta 1199 (1994) 105-114 ally polymerize under certain conditions seems to indicate that the preferred IgG binding sites have certain symmetry that can facilitate inter-molecular interactions that favor some long range order. We should also point out that electron microscopy also failed to differentiate IgG molecules from ferritin unequivocally in ferritin-IgG conjugates [41]. The existence of large adhesion force in solution on these specimens is not fully understood at present. However, a plausible explanation is that loosely attached molecules (beyond the first few layers) became associated with the tip after engagement. This explanation would be consistent with the case of imaging D N A in air, where the adhesion force has been attributed to tip contamination [8,12]. Unfortunately, cleaning the tip on carbon coated mica did not yield consistent results for ferritin-IgG specimens. But we did observe that for tips having a large adhesion force, vigorous washing with salt solutions (PBS and 1 mM EDTA, or 1 M NaC1) often yielded some improvement on both the adhesion force and the image quality, probably due to the removal of some of the contaminants by the solution. These results indicate that the condition of the tip is still the most important factor in AFM imaging of biological samples and is also the most difficult to control. 3.3. Immunoglobulin M (IgM)
Fig. 5. Images of ferritin-IgG conjugates with tips showing no adhesion force. In (a), ferritin-IgG conjugates are seen polymerized.This image was obtained in 12 mM NaCI, 0.3 mM KCI and 1 mM phosphate buffer, pH 7.0. In (b), loosely distributed ferritin-IgG conjugates were imaged in deionized water. The irregular shape and the larger size are in clear distinction from those images of ferritin and apoferritin (Fig. 1 and Fig. 4). Scale bars: 200 nm (horizontal) and 50 nm (vertical). Both samples were prepared by method 2 with 100 ~g/ml ferritin-IgG conjugates in PBS buffer. No filtering was applied.
molecules are seen to polymerize (for unknown reasons) on the mica surface and in Fig. 5(b), lower coverage revealed individual molecules, although they were rarely completely isolated. According to Sigma Chemicals, each ferritin should have, on average, 2.3 IgG molecules bound. But the A F M did not resolve each IgG molecules from the ferritin, probably due to the flexibility of the IgG and the shadowing effect of the ferritin (since the ferritin should be much more rigid with a larger height than the IgG). But the molecular profile is considerably different from both ferritin and apoferritin. The measured lateral size of ferritin-IgG conjugate is about 22 nm × 17 nm, consistent with the assumption of one ferritin with two IgG bound. The fact that ferritin-IgG conjugate can actu-
In view of the difficulty with ferritin-IgG conjugates, the presence of IgG molecules on the surface of ferritin apparently complicated the tip-sample interaction. Thus, we further tested the method with a pure antibody specimen - IgM. IgM consists of 5 IgG subunits and a J chain at its center, connected through disulfide bonds between neighboring IgGs and between the IgG and the J chain. The total molecular weight is between 900 and 1000 kDa [42]. The only direct structural elucidation of IgM is from electron microscopy which showed 5 dendrite like spikes radiating from the center [41,43]. The IgM proved to be even more difficult to image than ferritin-IgG conjugates. Although IgM readily adhered to the mica surface under various conditions, the tip-sample interaction is extremely complicated. The most notable phenomenon is the appearance of negative (or so-called inverted) contrast. Fig. 6 shows an example in which all molecules showed a negative contrast, although the specimen was quite stable with repeated scans of good reproducibility under the same tip. It was observed that for a tip that yielded negative contrast, the image occasionally reverted to positive contrast, but in this case stable imaging was often impossible. We believe that the negative contrast is the direct consequence of tip contamination, which was always accompanied by a measurable adhesion force.
J. Yang et aL / Biochimica et Biophysica Acta 1199 (1994) 105-I14
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Fig. 6. A negative contrast image of IgM, as observed when the tip
2 5 0 nm
had an adhesion force larger than 2 nN. The size of individual molecules is that of IgM (35 + 5 nm), but none of them showed any detail. With different tips/specimens, the molecular size of negative contrast images varied by a factor up to 2. This specimen was prepared by method 2 with an incubation temperature of 40°C (50 mg/mi IgM in PBS buffer). No filtering was applied.
Fig. 7. A typical IgM image obtained in deionized water at a scanning speed of 4.73 Hz and a probe force of 0.2 nN without adhesion force. The specimen was prepared by method 2 at an incubation temperature of ~ 45°C (20 m g / m l IgM in PBS). The specimen showed good protein coverage, with an average molecular size of 35 + 3 nm. The same specimen could show negative contrast for some tips with non-zero adhesion force. See text for more discussion. No filtering was applied.
For good tips with very small adhesion force, positive contrast images could be obtained on the same specimen that yielded negative contrast for other tips. Fig. 7 is a typical 'normal' image of IgM in deionized water as prepared by method 2. Fig. 8 shows a gallery of 10 typical IgM molecules selected from several spec-
imens. It is seen that both the five IgG subunits and the central ring are resolved. The variation in morphology is expected from such large flexible molecules. The
a
b
c
d
e
f
g
h
i
j
50 nm Fig. 8. A collection of 10 typical IgM images. The 5 IgG subunits are clearly recognizable, with finer details revealed in some of them. The variation in morphology may be the result of the flexibility of the molecules in solution which is consistent with electron microscopy results. Low pass filtering was applied to reduce high frequency noise.
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quality of these images are quite comparable to that of electron microscopy [41,43]. Measurements showed a molecular diameter of 35 + 3 nm, which is 17% larger than the size determined from electron microscopy, perhaps due to the incubation at 45°C which broke some of the disulfide bonds, resulting in a more extended molecular profile upon adsorption, although we cannot rule out the possibility that some tip broadening is involved. A n o t h e r possibility is that the dimension from electron microscopy is an underestimate due to possible shrinkage in negative staining [44]. Since no further purification was p e r f o r m e d , the smaller molecules seen in Fig. 7 could be the contaminants or the debris of IgM broken apart during incubation. The height measured from the A F M images is between 2 - 5 nm, which showed a relatively smaller compression. It is interesting to note that the height variation due to the probe force was also quite small for D N A and bacteriophage adsorbed on aminopropylytriethoxy silane treated mica when imaged in water. The reason for this p h e n o m e n o n is not understood [8]. One would normally expect the IgM to be more compressible than the ferritin. These images were reproducible and stable with fresh tips that showed positive contrast without adhesion force. These specimens could be stored at 4°C for more than 48 h. The A F M results of IgM once again indicated the importance of controlling the tip condition. An additional point raised from this experiment is that one must be careful in interpreting the A F M images, due to the existence of negative contrast. This is particularly a problem for closely packed specimens, where false results can easily arise from the negative contrast that may be difficult to recognize.
a
b
3 0 nm
3.4. Applications to several other macromolecules
Fig. 9. AFM images of IgA molecules obtained in deionized water with a probe force of 0.1 nN and a scanning speed of 4.73 Hz. Specimens were prepared by method 2 at an incubation temperature of 40°C (Protein concentration: 5 /zg/ml in PBS). Images shown were obtained after overnight storage at 4°C. (a) shows a large scale image where IgA molecules apparently polymerized.The double line structure should be the reflection of the IgG dimer in leA. The arrow indicates one such dimer that can be directly identified. (b) shows four examples of individual IgA molecules. Each monomer measures about 17 nmX 11 nm, consistent with the IgG dimension. Further details were not resolved at this resolution. Low pass filtering was applied to reduce high frequency noise.
To make sure that the method is indeed applicable for routine experiments, we have applied the method outlined in Sec. II to few m o r e macromolecules. Fig. 9 is the A F M image of I g A obtained in deionized water. The specimens were p r e p a r e d by method 2. IgA has been known to be a dimer of I g G molecules linked through a J chain [45]. Interestingly, IgA also polymerized on the mica surface (Fig. 9(a)), like the ferritin-IgG conjugates (Fig. 5(a)). However, the close association of IgA molecules makes it difficult for direct identification of each dimer in the image. F r o m these polymers, the I g A molecule has a normal size of 24 + 3 nm x 17 + 2 nm, consistent with the known values based on the I g G structure. On some areas, though, the dimer configuration could be clearly identified. Fig. 9(b) shows four such images. It is seen that each m o n o m e r has a size of ~ 17 x 11 nm and the relative position is somewhat variable, as one would expect. These values are consistent with an orientation in which the IgA
molecule is fully extended. Fig. 10 is an A F M image of phospholipid vesicles (DPPC) adsorbed on mica surface, p r e p a r e d by method 1. The vesicles were prepared by the method outlined by H u a n g [46], which should be primarily unilamella vesicles. At room temperature, D P P C should be in the gel phase (transition temperature: 41°C). These vesicles were surprisingly stable, with almost no adhesion force. If one would attribute protein adhesion to mica as electrostatic, since mica is negatively charged, these vesicles would not fit into the explanation, because phosphatidylcholine is neutral. One thing to notice is that the top of these vesicles was not flattened by the probe force, contrary to one might expect for these relatively large holo-structures (the average diameter of these vesicles is 48 nm, but the largest one has a diameter of 200 nm). This result seems in contradiction to that of ferritin (Fig. 1). Further study is needed to elucidated
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4/ Fig. 10. This is an AFM image of DPPC vesicles adsorbed on mica surface. The majority of the vesicles are smaller ones and their average diameter is 485:7 nm. The vesicles were prepared by the method described by Huang [46] with a buffer solution containing 140 mM NaCI, 10 mM Tris, pH 8.0. The specimen was prepared by method 1. The image was obtained in the same buffer with a probe force of 0.5 nN and a scanning speed of 4.73 Hz. It is noticed that the top of the vesicles, even the larger ones, is not flattened by the probe. No filtering was applied. Scale bars: 500 nm (horizontal) and 180 nm (vertical).
this phenomenon. If proteins are incorporated in these vesicles, it is highly likely that they can be resolved by AFM. Fig. 11 is the AFM image of hemoglobin, taken in deionized water. The specimen was prepared by method 2, but incubated at 4°C overnight for near monolayer coverage. With hemoglobin, we found that the proteins stuck to the tip very easily when the protein coverage on the substrate was more than one layer. For those good specimens, the image was very stable and reproducible, as long as the probe force was kept below 0.2 nN. The molecular dimension measured
Fig. 11. AFM image of hemoglobin on mica surface. The specimen was prepared by method 2 (5 /Lg/ml hemoglobin in PBS) incubated at 4°C overnight. The hemoglobin measures 6.5-I-0.9 nm, slightly larger than the result from X-ray diffraction [47], maybe an indication of some tip-related broadening. This image was obtained in deionized water with a probe force of ~ 0.1 nN and a scanning speed of 4.73 Hz. Low pass filtering was applied to reduce high frequency noise. Scale bars: 50 nm (horizontal) and 12 nm (vertical).
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is 6.5 + 0.9 nm, slightly larger than the actual size (5.0 nm × 5.5 nm × 6.4 nm) determined from X-ray diffraction [47]. These additional AFM images of various macromolecules convincingly demonstrated that the methods presented in Section 2 are indeed generally applicable to soluble macromolecules and mica surface is adequate for nm resolution atomic force microscopy of directly adsorbed macromolecules in aqueous solutions.
4. Concluding remarks Specimen preparation is perhaps the most important aspect of any imaging technique. This is particularly true for AFM, which was invented only a short few years ago. Only recently, the AFM for biological applications began to appear promising, as represented by the success with DNA and membrane proteins. In this study, it is demonstrated that, contrary to popular perception, mica is a remarkable substrate not only for membranes, but also for soluble proteins. The method of direct adsorption on mica surface yielded stable, reproducible and high resolution AFM images at nearly physiological conditions. Our data also indicated that the gross conformation of macromolecules was little changed by the surface interaction with mica, at least, at the resolution achieved. We are surprised at the finding that, the main problem of AFM of these soluble proteins is not the adhesion to the substrate. In fact, all soluble proteins we have tried, including albumin, IgG, hemocyanin and myosin, adhered well to the substrate and often too much rather than too little. The problem is then how to prepare a monolayer or just a few layers on a substrate (not necessarily mica) with good uniformity. An additional but very serious problem is how to prevent proteins from sticking to the tip, a factor we have very little control of at present. This may require the use of different tip materials. If indeed these materials can be found, we can then further experiment with the tip geometry in order to improve upon the resolution achieved so far. There is reason to believe that with a good monolayer coverage and a non-sticking tip, similar resolution as the cholera toxin (1-2 nm) should be achievable by this method. A minor discovery from these experiments is that many proteins polymerized upon adsorption on mica surface for unknown reasons. In principle, this could provide some indirect information about the conformation of these macromolecules. Considering the short history of biological AFM, with further refinement of both specimen preparation techniques and tip fabrication, the AFM should become a useful imaging instrument to supplement with other techniques. The AFM should be able to provide
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a resolution comparable to that of electron microscopy with a specimen preparation method not much more complicated than that for light microscopy.
Acknowledgements We would like to thank Drs. A.P. Somlyo, A.V. Somlyo and S.M. Lindsay for useful discussions. We would also like to thank Zhiyu Hu for technical assistance. Support from Whitaker Foundation, US Army Research Office, Jeffress Memorial Trust, National Science Foundation and National Institutes of Health is gratefully acknowledged.
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