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Resources How to convert PDF Flashcards into a Deck on Anki (self.Anki)

KnightOfOne 於 2 天前 * 發表

Hey everyone! So I spent yesterday trying to figure out how to convert a pdf file with flashcards into a deck on Anki. The majority of the information on the web is for individuals with coding experience or some level of a knowledge base in coding. Unfortunately, my knowledge base and understanding of coding was too minimal to use those sites and I had to learn. Therefore, I’m making this guide to help someone who might find themselves in my position.

Acknowledgements

So you have a pdf file you want to make into an Anki deck. You’ve combed through the web and stumbled upon Takenote but have no idea how to use it. That is the entire basis for this guide. Takenote I give all credit to Takenote’s creator and honestly thank them.

Pre-Guide Requirements

1. PDF file with flashcards with separate question and answer pages. Here's an example:

   Page 1

   Front 1	Front 2
   Front 3	Front 4

   Page 2

   Back 2	Back 1
   Back 4	Back 3

2. If possible, try to create a new pdf file with about 2 pages; so 1 set of the above mentioned pages. We'll use these to get the right height, widths, and top-left offset coordinates without running through your whole pdf. (This saves a lot of time and frustration).

3. You can still use this script if you have more then 4 cards per page.

Part 1. The Set-Up

1.1 Download a Linux based system

There isn’t a complete need for this since you can use the command prompt on Windows or Terminal on Mac OS. However, since you have little to no experience working with programming, I’d rather you not mess up your machine by incorrectly typing something. Also the exact commands I list will be for Linux, as the creator of Takenote primarily uses that.

Installing Linux Steps:

1. Download Virtual Box from VirtualBox and download the platform packages for whoever operating system you’re currently using.

2. Open and install the package.

3. Download Ubuntu Desktop from Ubuntu

4. Install Ubuntu under Virtual Box

...

In most modern chromatographic proce- dures, a mixture of substances to be fractionated is dissolved in a liquid (the “mobile” phase) and percolated through a column containing a porous solid matrix (the “stationary” phase). As solutes flow through the column, they interact with the stationary phase and are retarded. The retarding force depends on the properties of each solute. If the column is long enough, sub- stances with different rates of migration will be separated. The chromato- graphic procedures that are most useful for purifying proteins are classified according to the nature of the interaction between the protein and the sta- tionary phase.

In ion ex- change chromatography, charged molecules bind to oppositely charged groups that are chemically linked to a matrix such as cellulose or agarose. Anions bind to cationic groups on anion exchangers, and cations bind to an- ionic groups on cation exchangers. Perhaps the most frequently used anion exchanger is a matrix with attached diethylaminoethyl (DEAE) groups, and the most frequently used cation exchanger is a matrix bearing carboxymethyl (CM) groups. DEAE: Matrix¬CH 2 ¬CH 2 ¬NH(CH 2 CH 3 ) ⫹ 2 CM: Matrix¬CH 2 ¬COO ⫺ Proteins and other polyelectrolytes (polyionic polymers) that bear both positive and negative charges can bind to both cation and anion exchangers. The binding affinity of a particular protein depends on the presence of other ions that compete with the protein for binding to the ion exchanger and on the pH of the solution, which influences the net charge of the protein.

The column effluent is collected in a series of fractions. Proteins that bind tightly to the ion exchanger can be eluted (washed through the column) by applying a buffer, called the eluant, that has a higher salt concentration or a pH that reduces the affinity with which the matrix binds the protein.

In hydrophobic interaction chro- matography, the matrix material is lightly substituted with octyl or phenyl groups. At high salt concentrations, nonpolar groups on the surface of pro- teins “interact” with the hydrophobic groups; that is, both types of groups are excluded by the polar solvent (hydrophobic effects are augmented by increased ionic strength). The eluant is typically an aqueous buffer with decreasing salt concentrations, increasing concentrations of detergent (which disrupts hy- drophobic interactions), or changes in pH.

In gel filtration chromatography (also called size exclusion or molecular sieve chromatography), molecules are separated according to their size and shape. The stationary phase consists of gel beads containing pores that span a rela- tively narrow size range. The pore size is typically determined by the extent of cross-linking between the polymers of the gel material. If an aqueous solu- tion of molecules of various sizes is passed through a column containing such “molecular sieves,” the molecules that are too large to pass through the pores are excluded from the solvent volume inside the gel beads. These large mole- cules therefore traverse the column more rapidly than small molecules that pass through the pores (Fig. 5-7). Because the pore size in any gel varies to some degree, gel filtration can be used to separate a range of molecules; larger mol- ecules with access to fewer pores elute sooner (i.e., in a smaller volume of elu- ant) than smaller molecules that have access to more of the gel’s interior volume. Within the size range of molecules separated by a particular pore size, there is a linear relationship between the relative elution volume of a substance and the logarithm of its molecular mass (assuming the molecules have similar shapes). If a given gel filtration column is calibrated with several proteins of known molecular mass, the mass of an unknown protein can be conveniently estimated by its elution position

A striking characteristic of many proteins is their ability to bind specific molecules tightly but noncovalently. This property can be used to purify such proteins by affin- ity chromatography (Fig. 5-8). In this technique, a molecule (a ligand) that specifically binds to the protein of interest (e.g., a nonreactive analog of an enzyme’s substrate) is covalently attached to an inert matrix. When an impure protein solution is passed through this chromatographic material, the desired pro- tein binds to the immobilized ligand, whereas other substances are washed through the column with the buffer. The desired protein can then be recovered in highly purified form by changing the elution conditions to release the protein from the matrix. The great advantage of affinity chromatography is its ability to ex- ploit the desired protein’s unique biochemical properties rather than the small differences in physicochemical properties between proteins exploited by other chromatographic methods. Accordingly, the separation power of affinity chro- matography for a specific protein is often greater than that of other chromato- graphic techniques. Affinity chromatography columns can be constructed by chemically at- taching small molecules or proteins to a chromatographic matrix. In im- munoaffinity chromatography, an antibody is attached to the matrix in order to purify the protein against which the antibody was raised.

In metal chelate affinity chromatography, a divalent metal ion such as Zn 2⫹ or Ni 2⫹ is attached to the chromatographic matrix so that proteins bear- ing metal-chelating groups (e.g., multiple His side chains) can be retained. Recombinant DNA techniques (Section 3-5) can be used to append a segment of six consecutive His residues, known as a His tag, to the N- or C-terminus of the polypeptide to be isolated. This creates a metal ion–binding site that allows the recombinant protein to be purified by metal chelate chromatogra- phy. After the protein has been eluted, usually by altering the pH, the His tag can be removed by the action of a specific protease whose recognition sequence separates the (His) 6 sequence from the rest of the protein.

Electrophoresis, the migration of ions in an electric field, is described in Section 3-4B. Polyacrylamide gel electrophoresis (PAGE) of proteins is typ- ically carried out in polyacrylamide gels with a characteristic pore size, so the molecular separations are based on gel filtration (size and shape) as well as elec- trophoretic mobility (electric charge). However, electrophoresis differs from gel filtration in that the electrophoretic mobility of smaller molecules is greater than the mobility of larger molecules with the same charge density. The pH of the gel is high enough (usually about pH 9) so that nearly all proteins have net negative charges and move toward the positive electrode when the current is switched on. Molecules of similar size and charge move as a band through the gel. Following electrophoresis, the separated bands may be visualized in the gel by an appropriate technique, such as soaking the gel in a solution of a stain that binds tightly to proteins. If the proteins in a sample are radioactive, the gel can be dried and then clamped over a sheet of X-ray film. After a time, the film is developed and the resulting autoradiograph shows the positions of the radioactive components by a blackening of the film.

a protein of interest is available, it can be used to specifically detect the pro- tein on a gel in the presence of many other proteins, a process called immunoblotting or Western blotting that is similar to ELISA

In one form of polyacrylamide gel electrophoresis, the detergent sodium dodecyl sulfate (SDS) [CH 3 ¬(CH 2 ) 10 ¬CH 2 ¬O¬SO 3 ⫺ ]Na ⫹ is added to denature proteins. Amphiphilic molecules (Section 2-1C) such as SDS interfere with the hydrophobic interactions that normally stabilize pro- teins. Proteins assume a rodlike shape in the presence of SDS. Furthermore, most proteins bind SDS in a ratio of about 1.4 g SDS per gram protein (about one SDS molecule for every two amino acid residues). The large negative charge that the SDS imparts masks the proteins’ intrinsic charge. The net re- sult is that SDS-treated proteins have similar shapes and charge-to-mass ra- tios. SDS-PAGE therefore separates proteins purely by gel filtration effects, that is, according to molecular mass. Figure 5-9 shows examples of the resolving power and the reproducibility of SDS-PAGE. In SDS-PAGE, the relative mobilities of proteins vary approximately lin- early with the logarithm of their molecular masses (Fig. 5-10). Consequently, the molecular mass of a protein can be determined with about 5 to 10% ac- curacy by electrophoresing it together with several “marker” proteins of known molecular masses that bracket that of the protein of interest. Because SDS dis- rupts noncovalent interactions between polypeptides, SDS-PAGE yields the molecular masses of the subunits of multisubunit proteins. The possibility that subunits are linked by disulfide bonds can be tested by preparing samples for SDS-PAGE in the presence and absence of a reducing agent, such as 2- mercaptoethanol (HSCH 2 CH 2 OH), that breaks those bonds (Section 5-3A)

Although gel electrophoresis in its various forms is highly effective at separat- ing charged molecules, it can require up to several hours and is difficult to quantitate and automate. These disadvantages are largely overcome through the use of capillary electrophoresis (CE), a technique in which electrophore- sis is carried out in very thin capillary tubes (20- to 100- m inner diameter). Such narrow capillaries rapidly dissipate heat and hence permit the use of very high electric fields, which reduces separation times to a few minutes.

A protein has charged groups of both polarities and therefore has an isoelectric point, pI, at which it is immobile in an electric field. If a mixture of proteins is electrophoresed through a solution or gel that has a stable pH gradient in which the pH smoothly increases from anode to cathode, each protein will migrate to the po- sition in the pH gradient corresponding to its pI. If a protein molecule diffuses away from this position, its net charge will change as it moves into a region of different pH and the resulting electrophoretic forces will move it back to its iso- electric position. Each species of protein is thereby “focused” into a narrow band about its pI. This type of electrophoresis is called isoelectric focusing (IEF). IEF can be combined with SDS-PAGE in an extremely powerful separation technique named two-dimensional (2D) gel electrophoresis. First, a sample of proteins is subjected to IEF in one direction, and then the separated proteins are subjected to SDS-PAGE in the perpendicular direction. This procedure gen- erates an array of spots, each representing a protein (Fig. 5-11)

The rate at which a particle sediments in the ultracentrifuge is related to its mass (the density of the solution and the shape of the particle also affect the sedimentation rate).

Because a protein contains multiple charged groups, its solubility depends on the concentrations of dissolved salts, the polarity of the solvent, the pH, and the temperature. Some or all of these variables can be manipulated to selec- tively precipitate certain proteins while others remain soluble. The solubility of a protein at low ion concentrations increases as salt is added, a phenomenon called salting in. The additional ions shield the pro- tein’s multiple ionic charges, thereby weakening the attractive forces between individual protein molecules (such forces can lead to aggregation and precip- itation). However, as more salt is added, particularly with sulfate salts, the sol- ubility of the protein again decreases. This salting out effect is primarily a result of the competition between the added salt ions and the other dissolved solutes for molecules of solvent. At very high salt concentrations, so many of the added ions are solvated that there is significantly less bulk solvent avail- able to dissolve other substances, including proteins.

• To be sequenced, a protein must be separated into individual polypeptides that can be cleaved into sets of overlapping fragments. • The amino acid sequence can be determined by Edman degradation, a procedure for removing N-terminal residues one at a time. • Mass spectrometry can identify amino acid sequences from the mass-to-charge ratio of gas-phase protein fragments

The protein must be broken down into fragments small enough to be individually se- quenced, and the primary structure of the intact protein is then reconstructed from the sequences of overlapping fragments (Fig. 5-13). Such a procedure, as we have seen (Section 3-4C), is also used to sequence DNA.

Each polypeptide chain (if it is not chemically blocked) has an N-terminal residue. Identifying this “end group” can establish the number of chemically dis- tinct polypeptides in a protein. For example, insulin has equal amounts of the N-terminal residues Gly and Phe, which indicates that it has equal numbers of two nonidentical polypeptide chains.

Thus, it is possible to determine the amino acid sequence of a polypeptide chain from the N-terminus inward by sub- jecting the polypeptide to repeated cycles of Edman degradation and, after every cycle, identifying the newly liberated PTH-amino acid

Mass spectrometry has emerged as an important technique for characterizing and sequencing polypeptides. Mass spectrometry accurately measures the mass- to-charge (m兾z) ratio for ions in the gas phase (where m is the ion’s mass and z is its charge)

Newer techniques have addressed this shortcoming. For example, in the electrospray ionization (ESI) technique, a solution of a macromolecule such as a peptide is sprayed from a narrow capillary tube maintained at high volt- age (⬃4000 V), forming fine, highly charged droplets from which the solvent rapidly evaporates (Fig. 5-17a). This yields a series of gas-phase macromolec- ular ions that typically have ionic charges in the range ⫹0.5 to ⫹2 per kilo- dalton. The charges result from the protonation of basic side chains such as Arg and Lys. The ions are directed into the mass spectrometer, which meas- ures their m兾z values with an accuracy of ⬃0.01% (Fig. 5-17b). Consequently, determining an ion’s z permits its molecular mass to be determined with far greater accuracy than by any other method.

Homologous proteins with the same function in different species (e.g., the cytochromes c shown in Table 5-6) are said to be orthologous. Within a species, similar proteins arise through gene duplication, an aber- rant genetic recombination event in which one member of a chromosome pair acquires both copies of the primordial gene (genetic recombination is discussed in Section 25-6). Following duplication, the sequences may diverge as muta- tions occur over time. Gene duplication is a particularly efficient mode of evo- lution because one copy of the gene evolves a new function through natural selection while its counterpart continues to direct the synthesis of the original protein. Tw o independently evolving genes that are derived from a duplication event are said to be paralogous. In prokaryotes, approximately 60% of protein domains have been duplicated; in many eukaryotes the figure is ⬃90%, and in humans, ⬃98%

The globin family of proteins provides an excellent example of evolution through gene duplication and divergence. Hemoglobin, which transports O 2 from the lungs (or gills or skin) to the tissues, is a tetramer with the sub- unit composition ␣ 2 ␤ 2 (i.e., two ␣ polypeptides and two ␤ polypeptides). The sequences of the ␣ and ␤ subunits are similar to each other and to the se- quence of the protein myoglobin, which facilitates oxygen diffusion through muscle tissue (hemoglobin and myoglobin are discussed in more detail in Chapter 7). The primordial globin probably functioned simply as an oxygen- storage protein. Gene duplication allowed one globin to evolve into a monomeric hemoglobin ␣ chain. Duplication of the ␣ chain gene gave rise to the paralogous gene for the ␤ chain.

O montante de juros originário tende a ser diretamente proporcional à preferência temporal dos agentes econômicos, ou seja, quanto mais estes valorizem o consumo presente em relação ao futuro, maior deverá ser o montante de juros necessário para induzi-los a poupar, isto é, a postergar o consumo e, inversamente, quanto maior a preferência pelo consumo futuro em relação ao presente, menor deverá ser o total de juros que ele requererá para poupar. Se, por exemplo, os agentes econômicos soubessem que o fim do mundo seria no dia seguinte, a taxa de juros tenderia ao infinito; se, por outro lado, fossem informados de que nunca morreriam então a taxa de juros cairia para níveis baixíssimos, próximos de zero.

rates of evolution vary among proteins (Fig. 5-24). This does not imply that the rates of mutation of the DNAs specifying those proteins differ, but rather that the rate at which muta- tions are accepted into a protein varies. A major contributor to the rate at which a protein evolves is the effect of amino acid changes on the protein’s function. For example, Fig. 5-24 shows that histone H4,

The rate of protein evolution also depends on the protein’s structural sta- bility. For example, a mutation that slowed the rate at which a newly synthe- sized polypeptide chain folds into its functional three-dimensional shape could affect the cell’s survival, even if the protein ultimately functioned normally. Such mutations would be especially critical for proteins that are produced at high levels, since the not-yet-folded proteins could swamp the cell’s protein- folding mechanisms.

Mutational changes in proteins do not account for all evolutionary changes among organisms. The DNA sequences that control the expression of proteins (Chapters 26, 27, and 28) are also subject to mutation. These se- quences control where, when, and how much of the corresponding protein is made.

Finding the same residue at a particular position in the amino acid sequence of a series of related proteins suggests that the chemical or structural properties of that so-called invariant residue uniquely suit it to some essential function of the protein.

Other amino acid positions may have less stringent side chain re- quirements and can therefore accommodate residues with similar characteristics (e.g., Asp or Glu, Ser or Thr, etc.); such positions are said to be conservatively substituted. On the other hand, a particular amino acid position may tolerate many different amino acid residues, indicating that the functional requirements of that position are rather nonspecific. Such a po- sition is said to be hypervariable. Why is cytochrome c —an ancient and essential protein—not identical in all species? Even a protein that is well adapted to its function, that is, one that is not subject to physiological improvement, nevertheless continues evolving. The random nature of mutational processes will, in time, change such a protein in ways that do not significantly affect its function, a process called neutral drift (deleterious mutations are, of course, rapidly rejected through natural selection). Hypervariable residues are apparently particularly subject to neutral drift